Current Protocols in Stem Cell Biology

Current Protocols in Stem Cell Biology Online ISBN: 9780470151808 DOI: 10.1002/9780470151808

Table of Contents 1. Foreword 2. Preface 3. Chapter 1 Embryonic and Extraembryonic Stem Cells 1.

Section A Isolation of Embryonic Stem Cells 1. Introduction 2. Unit 1A.1 Derivation and Characterization of Nonhuman Primate Embryonic Stem Cells 3. Unit 1A.2 Derivation of hESC from Intact Blastocysts 4. Unit 1A.3 Reprogramming Primordial Germ Cells (PGC) to Embryonic Germ (EG) Cells 5. Unit 1A.4 Derivation and Propagation of hESC Under a Therapeutic Environment

2. Section B Characterization of Embryonic Stem Cells 1. Unit 1B.1 Proteomic Analysis of Pluripotent Stem Cells 2. Unit 1B.2 Gene Expression Analysis of RNA Purified from Embryonic Stem Cells and Embryoid Body–Derived Cells Using a High-Throughput Microarray Platform 3. Unit 1B.3 Phenotypic Analysis of Human Embryonic Stem Cells 4. Unit 1B.4 Isolation of Human Embryonic Stem Cell–Derived Teratomas for the Assessment of Pluripotency 5. Unit 1B.5 Tandem Affinity Purification of Protein Complexes in Mouse Embryonic Stem Cells Using In Vivo Biotinylation 6. Unit 1B.6 Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells 7. Unit 1B.7 Preparation of Defined Human Embryonic Stem Cell Populations for Transcriptional Profiling 3. Section C Culture and Maintenance of Undifferentiated Embryonic Stem Cells 1. Introduction 2. Unit 1C.1 Expansion of Human Embryonic Stem Cells In Vitro 3. Unit 1C.2 Defined, Feeder-Independent Medium for Human Embryonic Stem Cell Culture 4. Unit 1C.3 Isolation and Propagation of Mouse Embryonic Fibroblasts and Preparation of Mouse Embryonic Feeder Layer Cells 5. Unit 1C.4 Culture of Mouse Embryonic Stem Cells 6. Unit 1C.5 Preparation of Autogenic Human Feeder Cells for Growth of Human Embryonic Stem Cells 7. Unit 1C.6 Isolation of Human Placental Fibroblasts 8. Unit 1C.7 Derivation of Human Skin Fibroblast Lines for Feeder Cells of Human Embryonic Stem Cells 9. Unit 1C.8 Cryopreservation of Dissociated Human Embryonic Stem Cells in the Presence of ROCK Inhibitor 10. Unit 1C.9 Authentication and Banking of Human Pluripotent Stem Cells 11. Unit 1C.10 Clump Passaging and Expansion of Human Embryonic and Induced Pluripotent Stem Cells on Mouse Embryonic Fibroblast Feeder Cells 12. Unit 1C.11 Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers 4. Section D Germ Layer Induction/Differentiation of Embryonic Stem Cells 1. Unit 1D.1 Germ Layer Induction in ESC—Following the Vertebrate Roadmap 2. Unit 1D.2 Formation and Hematopoietic Differentiation of Human Embryoid Bodies by Suspension and Hanging Drop Cultures 3. Unit 1D.3 Directed Differentiation of Human Embryonic Stem Cells as Spin Embryoid Bodies and a Description of the Hematopoietic Blast Colony Forming Assay 4. Unit 1D.4 Differentiation of Human Embryonic Stem Cells in Adherent and in Chemically Defined Culture Conditions 5. Unit 1D.5 Isolation and Differentiation of Xenopus Animal Cap Cells 5. Section E Extraembryonic Lineages 1. Introduction 2. Unit 1E.1 Isolation of Human Placenta-Derived Multipotent Cells and In Vitro Differentiation into Hepatocyte-Like Cells 3. Unit 1E.2 Isolation of Mesenchymal Stem Cells from Amniotic Fluid and Placenta

4. 5. 6. 7.

Unit 1E.3 Isolation of Amniotic Epithelial Stem Cells Unit 1E.4 Isolation and Manipulation of Mouse Trophoblast Stem Cells Unit 1E.5 Isolation of Amniotic Mesenchymal Stem Cells Unit 1E.6 Amnion Epithelial Cell Isolation and Characterization for Clinical Use

6. Section F Mesodermal Lineages 1. Unit 1F.1 Differentiation of Embryonic Stem Cells into Cartilage Cells 2. Unit 1F.2 Differentiation of Human Embryonic Stem Cells to Cardiomyocytes by Coculture with Endoderm in Serum-Free Medium 3. Unit 1F.3 Isolation of Hematopoietic Stem Cells from Mouse Embryonic Stem Cells 4. Unit 1F.4 Differentiation of Mouse Embryonic Stem Cells into Blood 5. Unit 1F.5 Endothelial Differentiation of Embryonic Stem Cells 6. Unit 1F.6 Hematopoietic Differentiation of Human Embryonic Stem Cells by Cocultivation with Stromal Layers 7. Unit 1F.7 TLX1 (HOX11) Immortalization of Embryonic Stem Cell–Derived and Primary Murine Hematopoietic Progenitors 8. Unit 1F.8 Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts from Human Embryonic Stem Cells 9. Unit 1F.9 Derivation of Vasculature from Embryonic Stem Cells 10. Unit 1F.10 Isolation and Functional Characterization of Pluripotent Stem Cell–Derived Cardiac Progenitor Cells 11. Unit 1F.11 Differentiation of Mouse Embryonic Stem Cells into Cardiomyocytes via the Hanging-Drop and Mass Culture Methods 7. Section G Endodermal Lineages 1. Unit 1G.1 The Differentiation of Distal Lung Epithelium from Embryonic Stem Cells 2. Unit 1G.2 Pancreas Differentiation of Mouse ES Cells 3. Unit 1G.3 Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm 8. Section H Ectodermal Lineages 1. Unit 1H.1 Differentiation of Mouse Embryonic Stem Cells to Spinal Motor Neurons 2. Unit 1H.2 Time-Lapse Imaging of Embryonic Neural Stem Cell Division in Drosophila by Two-Photon Microscopy

4. Chapter 2 Somatic Stem Cells 1.

Section A Hematopoietic Stem Cells 1. Introduction 2. Unit 2A.1 Isolation of Mononuclear Cells from Human Cord Blood by Ficoll-Paque Density Gradient 3. Unit 2A.2 Isolation of Hematopoietic Stem Cells from Human Cord Blood 4. Unit 2A.3 Isolation of Mesenchymal Stem Cells from Human Cord Blood 5. Unit 2A.4 Isolation and Assessment of Long-Term Reconstituting Hematopoietic Stem Cells from Adult Mouse Bone Marrow 6. Unit 2A.5 Analysis of the Hematopoietic Stem Cell Niche 7. Unit 2A.6 Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues 8. Unit 2A.7 High Level In Vitro Expansion of Murine Hematopoietic Stem Cells 9. Unit 2A.8 Isolation and Visualization of Mouse Placental Hematopoietic Stem Cells 10. Unit 2A.9 Preparation of Hematopoietic Stem and Progenitor Cells from the Human Placenta

2. Section B Non-Hematopoietic Bone Marrow-Derived Stem Cells 1. Unit 2B.1 Isolation and Characterization of Mesoangioblasts from Mouse, Dog, and Human Tissues 2. Unit 2B.2 Purification and Culture of Human Blood Vessel–Associated Progenitor Cells 3. Unit 2B.3 Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells 3. Section C Cardiovascular Stem Cells 1. Unit 2C.1 Isolation and Characterization of Endothelial Progenitor Cells from Human Blood 2. Unit 2C.2 Derivation of Epicardium-Derived Progenitor Cells (EPDCs) from Adult Epicardium 3. Unit 2C.3 Isolation and Expansion of Cardiosphere-Derived Stem Cells 4. Section D Neural Stem Cells 1. Unit 2D.1 Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains 2. Unit 2D.2 Isolating, Expanding, and Infecting Human and Rodent Fetal Neural Progenitor Cells 3. Unit 2D.3 Long-Term Multilayer Adherent Network (MAN) Expansion, Maintenance, and Characterization, Chemical and Genetic Manipulation, and Transplantation of Human Fetal Forebrain Neural Stem Cells

4. 5. 6.

Unit 2D.4 Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation Unit 2D.5 Isolation and Culture of Ventral Mesencephalic Precursor Cells and Dopaminergic Neurons from Rodent Brains Unit 2D.6 Isolation of Neural Stem Cells from Neural Tissues Using the Neurosphere Technique

5. Section E Germline Stem Cells 1. Unit 2E.1 Culturing Ovarian Somatic and Germline Stem Cells of Drosophila 2. Unit 2E.2 Time-Lapse Live Imaging of Stem Cells in Drosophila Testis 6. Section F Gut Stem Cells 1. Unit 2F.1 In Situ Hybridization to Identify Gut Stem Cells 7. Section G Lung Stem Cells 1. Unit 2G.1 Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

5. Chapter 3 Cancer Stem Cells 1. Unit 3.1 Colon Cancer Stem Cells 2. Unit 3.2 In Vivo Evaluation of Leukemic Stem Cells through the Xenotransplantation Model 3. Unit 3.3 Culture and Isolation of Brain Tumor Initiating Cells

6. Chapter 4 Manipulation of Potency 1.

Section A iPS Cells 1. Unit 4A.1 Human iPS Cell Derivation/Reprogramming 2. Unit 4A.2 Generation and Characterization of Human Induced Pluripotent Stem Cells

2. Section B Nuclear Transfer 1. Unit 4B.1 Heterokaryon-Based Reprogramming for Pluripotency

7. Chapter 5 Genetic Manipulation of Stem Cells 1.

Section A Lineage Tracers in Stem Cells 1. Unit 5A.1 Imaging Neural Stem Cell Fate in Mouse Model of Glioma 2. Unit 5A.2 Functional Analysis of Adult Stem Cells Using Cre-Mediated Lineage Tracing 3. Unit 5A.3 Magnetic Resonance Imaging of Human Embryonic Stem Cells 4. Unit 5A.4 Lineage Tracing in the Intestinal Epithelium 5. Unit 5A.5 Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes in Mammalian Neocortical Neural Progenitors

2. Section B Homologous Recombination in Stem Cells 6. Unit 5B.1 Generation of Human Embryonic Stem Cell Reporter Knock-In Lines by Homologous Recombination

8. Appendix 1 Useful Information

1. 1A Guidelines for the Conduct of Human Embryonic Stem Cell Research 2. 1B ISSCR Guidelines for the Clinical Translation of Stem Cells

9. Appendix 2 Laboratory Equipment Standard Laboratory Equipment

10. Appendix Suppliers

Selected Suppliers of Reagents and Equipment

FOREWORD tem cell biology is emerging as a field in biology with tremendous therapeutic potential. Making this potential a reality requires an international effort. The recognition that such a promising yet multifaceted discipline needs fostering led to the establishment of the International Society for Stem Cell Research (ISSCR). Central to the efforts of the ISSCR is the development of tools to ensure the success of stem cell researchers. What better way to do this than to collaborate with Current Protocols to develop this valuable compendium of protocols in stem cell biology?

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Stem cell researchers have developed a number of breakthrough techniques, from the derivation and manipulation of pluripotent embryonic stem cells to purification and transplantation of tissue-restricted stem cells from adult organs. A number of laboratories have become leaders in the field as a result of developing such techniques. The more efficient scientists are at implementing new and powerful methodologies in their own laboratories, the faster stem cell biology will advance our understanding of normal development and lead to the development of therapies. Thus, the availability of quality protocols will have a major impact on the success of the entire field. Current Protocols has long been the premier volume for proven in-depth protocols regarding many aspects of biology, and this volume on stem cell biology will prove a valuable addition to researchers worldwide. Experiments in stem cell biology must be interpreted with great caution as well as openness to alternative explanations. For example, the recently discovered phenomenon of cell fusion in vivo or the existence of tissue-restricted blood stem cells in peripheral tissues were initially misinterpreted as evidence for stem cell trans-differentiation. It is very important that this compendium of protocols highlight potential pitfalls as well as maintain the opportunity for clarification and correction when the need arises. The fact that these protocols will be provided online will help ensure that researchers always have the latest, most up-to-date protocols available to them. The stem cell field is burgeoning, and, as I have seen within the ISSCR, there is a genuine push to share information and interact so that the field can move forward quickly. There is a drive to develop not only excellent basic research skills but to bring the findings to clinical use so that patients can benefit. As a Hematology Attending Physician at Children’s Hospital Boston, I treat children who have pediatric blood diseases or leukemia and am drawn by the need to translate our research findings into therapies to help treat a number of diseases. The promise is great, but we need to deliver, and I believe Current Protocols in Stem Cell Biology will help tremendously. Leonard I. Zon

Current Protocols in Stem Cell Biology Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.scfores1 C 2007 John Wiley & Sons, Inc. Copyright


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PREFACE he concept of tissue regeneration was already present in ancient Greece, reflected by the mythological stories of Prometheus or the Hydra, and described by Aristotle. The first scientific studies of the phenomenon were performed around 1740 by Abraham Trembley on the cnidarian polyp Hydra. Yet, it took another 150 years until the idea emerged that tissue maintenance, turnover, and regeneration may be rooted in rare cells with unique properties: stem cells.

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During the past 50 years, the development and improvement of techniques to isolate, track, manipulate, culture, characterize, and transplant cells has led to the discovery of stem cells in many different tissues. The power of the hematopoietic stem cell to repopulate the entire blood system, first demonstrated by Till and McCulloch in 1963, has long since been harnessed for clinical use. More recently, the identification of the even more versatile pluripotent embryonic stem cell by Evans and Kaufman (1981) and Martin (1981) has revolutionized our ability to probe mammalian developmental biology and to model human diseases. In recent years the fascination of scientists with stem cells has spilled over into the public domain, and many share the hope that the 21st century will see a revolution in regenerative medicine as novel therapies are derived from stem cells. Continued scientific study of the biology of stem cells will be critical for this prospect to become a reality. It is the goal of the editors, in developing this manual, to facilitate this endeavor by providing scientists with a compendium of well established protocols in stem cell biology. Along with the continued progress of the field of stem cell biology, this collection of protocols will expand. The manual is written such that even a seasoned stem cell biologist will find many novel and useful ideas, but with enough detail provided to also guide those with less experience. This product is not intended to substitute for a graduate course in stem cell biology or for a comprehensive textbook in the field. Introductory texts on stem cells and cell and developmental biology that we recommend include Handbook of Stem Cells (Lanza et al., 2004), Developmental Biology (Gilbert, 2006), and Molecular Cell Biology (Lodish et al., 2004) or Molecular Biology of the Cell (Alberts et al., 2002). We also strongly recommend that readers gain first-hand experience in basic laboratory techniques and safety procedures by training in a well established laboratory. Finally, with the great promise and potential of stem cells, come ethical concerns. We urge stem cell biologists to reflect on these issues and to respect internationally accepted ethical guidelines and limitations such as those developed by the International Society for Stem Cell Research on the Conduct of Human Embryonic Stem Cell Research (ISSCR; see APPENDIX A1.1).

HOW TO USE THIS MANUAL Format and Organization This publication is available online, with monthly supplements. Subjects in this manual are organized by chapters, which are subdivided into sections that contain protocols organized in units. Protocol units, which constitute the bulk of the title, generally describe a method and include one or more protocols with listings of materials, steps and annotations, recipes for unique reagents and solutions, and commentaries on the

Current Protocols in Stem Cell Biology iii-vi Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.scprefs1 C 2007 John Wiley & Sons, Inc. Copyright


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“hows” and “whys” of the method. Other units present more general information in the form of explanatory text with no protocols. Overview units contain theoretical discussions that lay the foundation for subsequent protocols, while discussion units present more general information. Page numbering in the PDF version reflects the modular arrangement by unit; for example, page 1A.2.3 refers to Chapter 1 (Embryonic and Extraembryonic Stem Cells), Section A (Isolation of Embryonic Stem Cells, UNIT 1.2 (Derivation of hESCs from Intact Blastocysts), and page 3 of that particular unit. Although many reagents and procedures are employed repeatedly throughout the manual, we have opted to retain individual authors’ recipes or supplier designations because of the importance of using a particular reagent or procedure for successful stem cell experiments. Cross-referencing among the units is used for very basic procedures that do not vary from laboratory to laboratory.

Introductory and Explanatory Information Because this publication is first and foremost a compilation of laboratory techniques in stem cell biology, we have included explanatory information where required to help readers gain an intuitive grasp of the procedures. Some sections begin with special overview units that describe the state of the art of the topic matter and provide a context for the procedures that follow. Section and unit introductions describe how the protocols that follow connect to one another, and annotations to the actual protocol steps describe what is happening as a procedure is carried out. Finally, the Commentary that closes each protocol unit describes background information regarding the historical and theoretical development of the method, as well as alternative approaches, critical parameters, troubleshooting guidelines, anticipated results, and time considerations. All units contain cited references and many indicate key references to inform users of particularly useful background reading, original descriptions, or applications of a technique. Protocols Many units in the manual contain groups of protocols, each presented with a series of steps. One or more basic protocols are presented first in each unit and generally cover the recommended or most universally applicable approaches. Alternate protocols are provided where different equipment or reagents can be employed to achieve similar ends, where the starting material requires a variation in approach, or where requirements for the end product differ from those in the basic protocol. Support protocols describe additional steps that are required to perform the basic or alternate protocols; these steps are separated from the core protocol because they might be applicable to other uses in the manual or because they are performed in a time frame separate from the basic protocol steps. Reagents and Solutions Reagents required for a protocol are itemized in the materials list before the procedure begins. Many are common stock solutions, others are commonly used buffers or media, while others are solutions unique to a particular protocol. Recipes for solutions are provided in each unit, following the protocols (and before the commentary) under the heading Reagents and Solutions. It is important to note that the names of some of these special solutions might be similar from unit to unit (e.g., RIPA buffer) while the recipes differ; thus, make certain that reagents are prepared from the proper recipes.

Preface

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Commercial Suppliers Throughout the manual, the authors have recommended commercial suppliers of chemicals, biological materials, and equipment. It is recommended that the user follow the author’s designations; often those are the products that the author, after considerable experimentation, has found will work under the particular conditions. In other cases, the experience of the author of that protocol is limited to that brand. In the latter situation, recommendations are offered as an aid to the novice in obtaining the tools of the trade. Phone numbers, facsimile numbers, and URLs of all suppliers mentioned in this manual are provided in the SUPPLIERS APPENDIX.

Safety Considerations Anyone carrying out these protocols may encounter the following hazardous or potentially hazardous materials: (1) radioactive substances, (2) toxic chemicals and carcinogenic or teratogenic reagents, and (3) pathogenic and infectious biological agents. Check the guidelines of your particular institution with regard to use and disposal of these hazardous materials. Although cautionary statements are included in the appropriate units, we emphasize that users must proceed with the prudence and precaution associated with good laboratory practice, and that all materials must be used in strict accordance with local and national regulations. Animal Handling Many protocols call for use of live animals (usually rats or mice) for experiments. Prior to conducting any laboratory procedures with live subjects, the experimental approach must be submitted in writing to the appropriate Institutional Animal Care and Use Committee (IACUC) or must conform to appropriate governmental regulations regarding the care and use of laboratory animals. Written approval from the IACUC (or equivalent) committee is absolutely required prior to undertaking any live-animal studies. Some specific animal care and handling guidelines are provided in the protocols where live subjects are used, but check with your IACUC or governmental guidelines to obtain more extensive information. Human Material See the International Society for Stem Cell Research “Guidelines for the Conduct of Human Embryonic Stem Cell Research,” reproduced in APPENDIX A1.1. Research using human tissues must be reviewed and approved by the independent institutional ethics review panel, and donated material must be provided voluntarily with informed consent. Reader Response Most of the protocols included in this manual are used routinely in the authors’ laboratories. These protocols work for them; to make them work for you the authors have annotated critical steps and included critical parameters and troubleshooting guides in the commentaries to most units. However, the successful evolution of this manual depends upon readers’ observations and suggestions. Consequently, we encourage readers to send in their comments ([email protected]). Current Protocols in Stem Cell Biology

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ACKNOWLEDGMENTS This manual is the product of dedicated efforts by many of our scientific colleagues who are acknowledged in each unit and by the hard work by the Current Protocols editorial staff at John Wiley and Sons. We are extremely grateful for the critical contributions by Kathy Morgan (Series Editor), who kept the editors and the contributors on track and played a key role in bringing the entire project into existence. Other skilled members of the Current Protocols staff who contributed to the project include Joseph White, Tom Cannon, and Sheila Kaminsky. The extensive copyediting required to produce an accurate protocols manual was ably handled by Allen Ranz, Susan Lieberman, Marianne Huntley, and Sylvia de Hombre. Typesetting and electronic illustrations were prepared by Aptara.

RECOMMENDED BACKGROUND READING Alberts, B., Roberts, K., Lewis, J., Raff, M., Walter, P., and Johnson, A. 2002. Molecular Biology of the Cell, 2nd ed. Garland Publishing, New York. Gilbert, S. 2006. Developmental Biology, 8th ed. Sinauer Publishing, Sunderland, Mass. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Lanza, R., Weissman, I., Thomson, J., Pedersen, R., Hogan, B., Gearhart, J., Blau, H., Melton, D., Moore, M., Verfailllie, C., Donnall Thomas, E., and West, M. (eds.) 2004. Handbook of Stem Cells. Elsevier, New York. Lodish, H., Berk, A., Matsudaira, P., Kaiseer, C.A., Krieger, M., Scott, M.P., Zipursky, L., and Darnell, J. 2004. Molecular Cell Biology, 5th ed. W.H. Freeman and Company, New York. Martin, G.R. 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. 78:7634-7638. Till, J.E. and McCulloch, E.A. 1963. Early repair processes in marrow cells irradiating and proliferating in vivo. Radiat. Res. Jan. 18:96-105.

Mick Bhatia, Andrew Elefanty, Susan J. Fisher, Roger Patient, Thorsten M. Schlaeger, and Evan Y. Snyder

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SECTION 1A Isolation of Embryonic Stem Cells INTRODUCTION erivation of first primate and soon thereafter human embryonic stem cells set the stage for the next exciting chapters in the stem cell field, in which we are beginning to learn the extent to which lessons learned from studying model systems apply to primate species. The commonalities will certainly be easier to discern than the unique aspects. However, before either is apparent, investigators need access to high-quality primate embryonic stem cell lines that are the truest in vitro representatives of their in vivo counterparts. In the case of mouse embryonic stem cells, it took decades for the field to establish criteria for their evaluation and produce lines that met them. In the context of work on model systems, it is virtually certain that many more primate embryonic stem cell lines must be produced before we know that we have the tools needed to delve deeper into major questions regarding the cells’ capacity for self-renewal as well as for differentiation.

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For these reasons this section includes detailed protocols for producing embryonic stem cells from both nonhuman primates (UNIT 1A.1) as well as humans (UNIT 1A.2). Not surprisingly, the methods are not dramatically different. However, special considerations apply in each case. For example, in nonhuman primates, complement-mediated lysis of the trophectoderm layer is deemed preferable to remove these cells before the stem cell derivation process begins. In contrast, many investigators who are producing new human embryonic stem cell lines wish to avoid their exposure to animal products such as antibodies. Thus, they opt to use intact embryos for derivation purposes and allow the trophectoderm layer to die during generation of the stem cell lines. Eventually, we will want to know if the presence or absence of trophoblasts, which contribute to the placenta, is a positive, negative, or neutral factor with regard to influencing embryonic stem cells quality. It is interesting to note from the numerous details that both groups include in their protocols, the complexity of the derivation process and the commitment this work requires. It takes a great deal of expertise to grow and manipulate human and nonhuman primate embryos. It requires vigilant monitoring of the cultures as the initial outgrowths form. A crucial step is making decisions about when the cultures should be divided. Although the authors have attempted to give as much specific information as possible about these steps, qualitative aspects of decision making remain that are subject to individual judgments best made based on experience. Finally, we note that the protocols focus on laboratory methods rather than ethical considerations, such as how to properly describe these studies to institutional review boards and how to obtain informed consent from donors. The enormity of these issues, which are handled in different ways by different institutions, are beyond the scope of this section but are of primary consideration to all investigators who are involved in both the derivation and use of new human and nonhuman primate embryonic stem cell lines. Susan J. Fisher Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1A.0.1 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01a00s1 C 2007 John Wiley & Sons, Inc. Copyright


1A.0.1 Supplement 1

Derivation and Characterization of Nonhuman Primate Embryonic Stem Cells

UNIT 1A.1

Christopher S. Navara,1 Carrie Redinger,1 Jocelyn Mich-Basso,1 Stacie Oliver,1 Ahmi Ben-Yehudah,1 Carlos Castro,1 and Calvin Simerly1 1

University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania

ABSTRACT Embryonic stem (ES) cells are a powerful research tool enabling the generation of mice with custom genetics, the study of the earliest stages of mammalian differentiation in vitro and, with the isolation of human ES cells, the potential of cell based therapies to a number of diseases including Parkinson’s and Type 1 diabetes. ES cells isolated from non-human primates offer the opportunity to ethically test the developmental potential of primate ES cells in chimeric offspring. If these cells have similar potency to mouse ES cells we may open a new era of primate models of human disease. Non-human primates are the perfect model system for the preclinical testing of ES cell–derived therapies. In this unit we describe methods for the derivation and characterization of non-human primate ES cells. With these protocols the investigator will be able to isolate nhpES cells and perform the necessary tests to confirm the pluripotent phenotype. Curr. Protoc. Stem C 2007 by John Wiley & Sons, Inc. Cell Biol. 1:1A.1.1-1A.1.21.
Keywords: nonhuman primate r embryonic stem cells r Oct-4 r Nanog r karyotype r teratoma

INTRODUCTION The use of murine embryonic stem (mES) cells has revolutionized the production of transgenic knockout, knockin, and knockdown mice, and has furthered biomedical research perhaps more than any other technological advance. Murine embryonic stem cells are stably growing cell lines that retain the ability to be recombined with cleavagestage embryos to produce animals with tissues derived from both the embryo and the stem cells. Alternatively, in a very elegant experimental procedure, embryonic stem cells can be combined with an experimentally derived tetraploid embryo. Tetraploid mouse embryos only form trophectoderm and extra-embryonic tissues during development. In these experiments, the resulting animal, including the germ line, is completely derived from the embryonic stem cells (Maatman et al., 2003). The overriding superiority of this technology is that transfection can be carried out on the mES cells using highly efficient techniques optimized for cultured cell lines. Selection of expression characteristics and stability of the transgene can be analyzed in vitro prior to generating transgenic animals. As the embryonic stem cells can be propagated, a large number of transgenic animals can be made in the F1 generation. Human embryonic stem cells (hESC), first isolated in 1998 (Thomson et al., 1998), hold great promise for cell-mediated therapies for debilitating diseases such as diabetes and Parkinson’s disease. These cells appear to be immortal in culture and retain the ability to form all tissues of the adult even through more than 100 passages. Due to obvious ethical concerns, the ability of these cells to contribute to chimeric offspring and the germ line has not, and should not, be tested; consequently, it is unknown if these cells share that important developmental property with mouse embryonic stem cells.

Current Protocols in Stem Cell Biology 1A.1.1-1A.1.21 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01a01s1 C 2007 John Wiley & Sons, Inc. Copyright


Isolation of Embryonic Stem Cells

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Nonhuman primate embryonic stem cells (nhpESC) were first isolated in 1995 from in vivo–fertilized rhesus embryos (Macaca mulatta; Thomson et al., 1995) and in 1996 from marmosets (Callithrix jacchus; Thomson et al., 1996). They have also been isolated from in vitro–fertilized (IVF) and intracytoplasmic sperm injection (ICSI)–fertilized (Suemori et al., 2001) and parthenogenetic cynomolgus monkeys (Cibelli et al., 2002; Vrana et al., 2003). These cells may prove invaluable in several ways. First, they serve as a preclinical model for testing the efficacy and safety of embryonic stem cell–derived therapies (Sanchez-Pernaute et al., 2005; Takagi et al., 2005). Secondly, they may enable the generation of nonhuman primates (NHP) expressing disease conditions as preclinical models of human disease. Some contribution of nhpESC to chimeric embryos has been shown, but no chimeric offspring have been generated (Takada et al., 2002; Mitalipov et al., 2006) to date. It is well established with regard to murine embryonic stem cells that some lines are able to contribute to fetal tissues but are deficient in their ability to contribute to the germ line. Therefore demonstrating that nhpES cells have this ability may require the derivation and testing of dozens of embryonic stem cell lines. In this unit, protocols are described for the high-efficiency derivation of embryonic stem cells from rhesus monkey embryos (Basic Protocol) and for the characterization of the pluripotent phenotype using immunocytochemistry (Support Protocol 1), RT-PCR (Support Protocol 2), and teratoma formation (Support Protocol 4). Additionally, as the generation of aneuploid cell lines is a recurring problem, a protocol is included for karyotyping nonhuman primate ES cells (Support Protocol 3). NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL

DERIVING NONHUMAN PRIMATE EMBRYONIC STEM CELLS Two basic techniques have been used for the isolation of embryonic stem cells. The first, described below, involves removing the outer trophectodermal cells of the expanded blastocyst using an antibody/complement reaction (“immunosurgery”). The tight junctions between trophectodermal cells prevent diffusion of the antibody into the inner cell mass (ICM), ensuring that only the trophectodermal cells bind antibody, and thus that they are the only cells lysed by the addition of complement. An alternative technique involves direct plating of the blastocyst without removal of the trophectoderm. This procedure also works successfully, but requires the investigator to later passage the inner cell mass (ICM) cells away from the trophectodermal cells in vitro. The former technique is included in this unit because it results in a cleaner embryonic stem cell preparation.

Materials

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

0.1% (w/v) gelatin in DPBS (Invitrogen, cat. no. 14190-144) Inactivated mouse embryonic fibroblasts (MEFs; Specialty Media, http://www.specialtymedia.com; also available from ATCC, cat. no. SCRC-1040.2) MEF medium (see recipe) nhpES cell medium (see recipe) Expanded non-human primate blastocysts (Hewitson, 2004) Acidified Tyrode’s medium (Chemicon) TALP-HEPES medium (see recipe) Anti-monkey serum produced in rabbit (Sigma, cat. no. M-0278) Mineral or silicon oil, embryo quality (Cooper Medical) Guinea pig complement, lyophilized (Biomeda; store at –20◦ C until use) Embryo-quality H2 O (Sigma)

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Current Protocols in Stem Cell Biology

Fetal bovine serum (FBS; Invitrogen, cat. no. 16000-044) Dimethylsulfoxide (DMSO) Liquid nitrogen 6-well tissue culture plate Hamilton syringe with 20-µl Unopette tip (Becton Dickinson) attached 37◦ C slide warmer 30-mm organ culture dish (Fisher) Dissecting microscope 60-mm non–tissue culture–treated petri dishes Stripping pipet: “Stripper” pipetting instrument (Fig. 1A.2.6) and 125-µm inner diameter plastic tips (MidAtlantic Diagnostics, http://www.midatlanticdiagnostics.com; cat. no. MXL3-125) Fine glass needle for passaging ES cells: pull a Pasteur pipet as thinly as possible while heating over Bunsen burner, such that a pair of needles with pointed sealed ends (mandatory) are produced, and bend according to preference for optimal access to the wells; alternatively, use commercially available stem cell knives (Swemed cat. no. 25111-109M; http://www.swemed.com) Cell scrapers 15-ml conical centrifuge tubes 1-ml cryovials Mr. Frosty freezing containers (Fisher) Prepare MEF plates 1. At a time point 48 hr prior to immunosurgery, prepare a gelatin-coated 6-well plate by placing 3 ml of 0.1% gelatin in PBS into each well and incubating in a sterile environment 1 to 2 hr at room temperature. 2. Rinse wells with MEF medium and plate 150,000 mitotically inactivated MEF cells/cm2 in 3 ml MEF medium. Return cells to incubator. The authors purchase MEFs from Specialty Media, but they are also available from ATCC; protocols exist for preparing them in one’s own laboratory, as well (Schatten et al., 2005). Plates containing MEFs are ready to use 24 to 48 hr after plating and should be used within 5 days. It is best to test MEFs before use, by culturing existing embryonic stem cell lines to determine that they support pluripotency

3. The day of the immunosurgery, remove the MEF medium and rinse each well with 2 ml nhpES cell medium. Discard rinse and add 3 ml of nhpES cell medium to each well. Return cells to incubator. This step should be performed well in advance of the immunosurgery (∼1 hr before), so that the medium is completely equilibrated before ICM plating.

Perform immunosurgery Embryos are always transferred using a Hamilton syringe with a 20-µl Unopette attached to the end. Monkey tissues are BSL-2 and should not be pipetted by mouth, as is common with mouse tissues. All immunosurgery steps are performed at 37◦ C on a prewarmed slide warmer. 4. Transfer rhesus expanded blastocysts (see Fig. 1A.1.1A) to 1 ml acidified Tyrode’s medium in a 30-mm organ culture dish. Observe under a dissecting microscope until the zona pellucida is removed (also see UNIT 1A.2). In very expanded blastocysts the zona is observed as a smooth, shiny region surrounding the embryo (Fig. 1A.1.1A); when it is successfully removed, the trophectoderm will become much more cellular.

5. Immediately after zona removal, transfer blastocysts into 3 to 5 ml TALP-HEPES medium and let stand for 5 min to wash. Current Protocols in Stem Cell Biology

Isolation of Embryonic Stem Cells

1A.1.3 Supplement 1

Figure 1A.1.1 Embryo to embryonic stem cells. (A) Nonhuman primate blastocysts should be fully expanded with a large distinct inner cell mass (ICM) prior to use for embryonic stem cell derivation. (B) After the complement is added the trophectoderm, cells are lysed, and the blastocyst will collapse. Lysed trophectodermal cells are only loosely associated with the ICM. (C) Isolated inner cell mass plated onto mouse embryonic feeder cells. Early passage nhpES cells for passaging should have very tightly packed cells with prominent nucleoli.

6. Prepare a 1:3 dilution of anti-monkey serum in TALP-HEPES medium (for a final concentration of 25% anti-monkey serum). In a 60-mm non–tissue culture–treated petri dish, place five to ten 10-µl drops of the diluted anti-monkey serum (number of drops will depend on number of blastocysts to be processed), then add just enough embryo-quality mineral or silicon oil to completely cover the drops. Warm to 37◦ C. 7. Transfer zona-free blastocysts to the drops of diluted anti-monkey serum and incubate on a 37◦ C slide warmer for 15 min. 8. Resuspend lyophilized guinea pig complement in 10 ml 4◦ C embryo-quality water, then prepare a 1:3 dilution of the reconstituted complement in TALP-HEPES medium (for a final concentration of 25% reconstituted complement) and keep on ice. Just prior to use, warm to 37◦ C and place 1 ml in an organ culture dish. Transfer embryos from anti-monkey serum directly into the complement solution. Incubate in the complement 15 min at 37◦ C. 9. Prepare a petri dish containing 50-µl drops of nhpES medium under oil, using the technique described in step 6. Briefly rinse the blastocysts with TALP-HEPES medium using the technique described in step 5 (but let stand only ∼30 sec instead of 5 min), transfer the blastocysts to the 50-µl drops of medium under oil, and return the blastocysts to the incubator for 30 min. The success of immunosurgery depends heavily on the blastocyst. Using the exact same conditions described here, the authors have observed classic lysis of the trophectodermal cells (Fig 1A.1.1B) and also almost no lysis of the trophectoderm. ES cells were successfully derived from both kinds of blastocysts. The dilution factor above applies to the lyophilized complement from Biomeda; other formulations may require different dilutions.

10. Draw the blastocyst into a stripping pipet with an inner diameter of 125 µm to remove the lysed cells and plate immediately (step 10). The diameter of the pipet is big enough to let the inner cell mass through but will strip the lysed cells from the ICM.

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

1A.1.4 Supplement 1

Plate isolated ICM on MEFs 11. Add 3 ml of nhpES cell medium to each well of a 6-well tissue culture dish containing an MEF feeder layer (prepared as in steps 1 to 3, above). Plate one isolated ICM (from step 9) or embryo (in case of failed immunosurgery or isolation without immunosurgery) into each well of the dish using the Hamilton syringe with 20-µl Unopette. The 6-well plate should only be opened in a biological safety cabinet. Current Protocols in Stem Cell Biology

12. Return plated embryos to the incubator. Do not disturb for at least 24 hr and preferably 48 hr. 13. After 48 hr, check the wells to determine if the ICM has firmly attached to the substrate (Fig. 1A.1.1C). If the ICM is firmly attached, replace 80% of the medium with 3 ml nhpES cell medium that has been preincubated at least 1 hr in a 37◦ C, 5% CO2 incubator in order to equilibrate it with the gas mixture and prewarm it to 37◦ C. If the ICM has not yet attached, add 3 ml of nhpES cell medium to the well. It is helpful at this point to use an objective marker to circle the location of the plated ICM.

14. Every 48 hr replace the medium with fresh nhpES cell medium. Continue for ∼14 days, until it becomes necessary to perform the first passage of the putative cell line. The cells should not be carried past 14 days without passaging, because the quality of the feeder cells will diminish and it is important to transfer the ES cells to fresh feeder cells. At this stage, cells that are promising will have a very large cell mass that may or may not look like embryonic stem cells. Prior to passaging on day 14, cell masses should be carefully watched for signs of retraction from the feeder layer. If this is observed, cell masses should be passaged immediately.

15. Manually passage any cells with proper embryonic stem cell morphology (high nuclei/cytoplasm ratio and prominent nucleoli; Fig. 1A.1.1D), cutting the cell masses into pieces containing 10 to 15 cells with a fine glass needle, and transferring the pieces to newly plated MEFs in a 6-well plate, as described above. Also cut and passage cell masses that do not resemble embryonic stem cells, if possible. If it is not possible to cut them manually, cell masses should be treated with 1 ml of 1 mg/ml collagenase and passaged. The authors have derived several stem cell lines from cell masses that did not initially have canonical embryonic stem cell morphology, so it is also advisable to attempt to culture these.

16. Maintain the initial culture plates for at least 1 week, changing medium every 48 hr, to determine if any other nhpES cell colonies begin to grow. 17. After the initial passage, passage cell lines approximately weekly using manual passaging, being sure to select only cells with proper ES cell morphology.

Freeze cells As soon as cultures are large enough to be split into three wells (day 6 or 7 after mechanical passaging), one well should be frozen. 18. Remove 6-well plate from incubator. Using a cell scraper, gently release ESCs and feeder layer from the bottom of the well. 19. Aspirate the cell suspension and place in a 15-ml conical tube. Rinse the well with 3 ml nhpES cell medium to resuspend any remaining cells and transfer to the same 15-ml tube. 20. Centrifuge 5 min at 200 × g, room temperature. During the centrifugation, prepare the freezing medium (90% v/v FBS containing 10% v/v DMSO). 21. When centrifugation is complete, remove and discard the supernatant. Resuspend pellet in 1 ml freezing medium. 22. Transfer resuspended cells to 1-ml cryovial. Place cryovial into a Mr. Frosty freezing container at room temperature. Place the Mr. Frosty freezing container in a –80◦ C freezer for 24 hr, then transfer to liquid nitrogen.

Isolation of Embryonic Stem Cells

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Supplement 1

Confirm and characterize the pluripotent phenotype 23. Once putative embryonic stem cells have been isolated, characterize them for the pluripotency markers SSEA-4, TRA160, and TRA 181 (Support Protocols 1 and/or 2), stable correct karyotype (Support Protocol 3), and ability to differentiate into tissues from all three germ layers (Support Protocol 4). The authors traditionally prefer to use immunocytochemistry (Support Protocol 1) whenever possible, as this allows for determining the heterogeneity of stem cell colonies. They also use RT-PCR (Support Protocol 2) to identify expression of genes related to pluripotency. The final criterion of pluripotency is the ability to form tissues derived from all three germ layers (Support Protocol 4). For determining this, the authors prefer teratoma formation, which offers straightforward technique and clear interpretation. Normal and stable karyotype is an important consideration when first deriving nonhuman primate ES cells, and is an ongoing concern while maintaining them. Included in this unit is a protocol for karyotyping nonhuman primate ES cells (Support Protocol 3). SUPPORT PROTOCOL 1

IMMUNOCYTOCHEMISTRY OF nhpES CELLS Immunocytochemistry has the advantage of measuring not only expression of a given protein but also the localization of the protein within the cell and within the embryonic stem cell colony. A number of pluripotency markers have been described for human and nonhuman primate embryonic stem cells. The classic markers are the transcription factors Nanog and Oct-4 and the surface antigen stage–specific embryonic antigen 3/4 (SSEA3/4), tumor rejection antigen (TRA) 1-60, and TRA 1-81. Primate embryonic stem cells are negative for the mouse embryonic stem cell marker SSEA1. Nanog and Oct-4 have functional relationships with pluripotency, whereas SSEA3/4, TRA 1-60, and TRA 1-81 are simply surface markers without a known function in embryonic stem cells.

Materials

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

1A.1.6 Supplement 1

0.1% (w/v) gelatin in DPBS Inactivated mouse embryonic fibroblasts (MEFs; Specialty Media, http://www.specialtymedia.com; also available from ATCC, cat. no. SCRC-1040.2) MEF medium (see recipe) Rhesus ES cells growing in culture (see Basic Protocol) Dulbecco’s phosphate-buffered saline (DPBS, Ca2+ - and Mg2+ -free; Invitrogen, cat. no. 14190-144), prewarmed DPBS containing 2% (v/v) formaldehyde DPBS containing 0.1% (v/v) Triton X-100 DPBS containing 0.3% (w/v) nonfat dry milk and 5% (v/v) normal goat serum Primary antibodies against desired ES markers (perform all dilutions in DPBS containing 0.1% v/v Triton X-100): Mouse Oct-4 [Santa Cruz Biotechnology (sc-5276); use at 1:100 dilution] Goat Nanog (R&D Systems; use at 1:20 dilution) Mouse SSEA-4 (Developmental Studies Hybridoma Bank; use at 1:5 dilution) Mouse TRA-1-81 (Santa Cruz Biotechnology; use at 1:5 dilution) Mouse TRA-1-60 (Santa Cruz Biotechnology; use at 1:5 dilution) Secondary antibody against IgG of species in which primary antibody was raised, labeled with Alexa Fluor 488; use at 100:1 dilution in DPBS containing 0.1% Triton X-100 10 mg/ml RNase in DPBS containing 0.1% Triton X-100 5 µM TOTO-3 (Invitrogen) in DPBS containing 0.1% Triton X-100 Vectashield mounting medium (Vector) Thermanox plastic coverslips (Ted Pella, Inc.) 6-well tissue culture plate Humidified chamber (e.g., Tupperware box containing moistened paper towels) Microscope slides Current Protocols in Stem Cell Biology

Prepare ES cells on MEF feeder layers for immunostaining 1. Prepare gelatin-coated Thermanox coverslips in a 6-well tissue culture plate containing one coverslip per well by placing 3 ml of 0.1% gelatin on the correct surface of each coverslip and incubating in a sterile environment 1 to 2 hr at room temperature. These coverslips are “sided”; medium will bead on the incorrect side.

2. Rinse coverslips with MEF medium and plate 15,625 MEF cells/cm2 on the gelatincoated surface. Incubate for 48 hr. 3. To prepare cells for immunostaining, passage nhpES cells (as described in Basic Protocol 1, step 15) onto the gelatin-coated Thermanox plastic coverslips seeded with MEF feeder cells and incubate ∼1 week prior to fixation and processing. Passaging and culture of cells is done as in the Basic Protocol, steps 15 to 17, except that in this protocol the wells of the 6-well plate contain coverslips.

Fix cells and block nonspecific binding 4. Prior to fixation, rinse coverslips with 3 ml warm DPBS to remove proteins found in the culture medium. 5. Transfer the coverslip immediately to 5 ml DPBS/2% formaldehyde and fix by incubating 40 min at room temperature. 6. After fixation, rinse cells with 5 to 7 ml DPBS/0.1% Triton X-100. 7. If necessary, block nonspecific binding of the antibodies at this stage using a 20-min incubation in 5 to 7 ml DPBS/0.3% (w/v) nonfat dry milk/5% (v/v) normal goat serum. Note that the Nanog antibody from R&D Systems is raised in goats, and blocking in goat serum will result in undesirable generalized staining masking the Nanog signal.

Treat cells with primary and secondary antibodies 8. Incubate sample coverslip with 100 µl primary antibody against the ES cell markers of interest at the appropriate dilution in DPBS/0.1% Triton X-100 for 40 min (except for Oct-4 and Nanog, which are most successfully stained at 4◦ C overnight) at 37◦ C in a humidified chamber. Alternative antibodies may work and investigators should determine their own optimal dilution.

9. Wash all samples for 15 min with DPBS/0.1% Triton X-100. 10. Add 100 µl fluorescently labeled secondary antibody to the sample coverslip and incubate for 40 min at 37◦ C in a humidified chamber. 11. Wash secondary antibody–exposed coverslip as described in step 9.

Counterstain and mount 12. Pretreat coverslip with 100 µl of 10 mg/ml RNase for 20 min. TOTO-3 will bind both RNA and DNA, so the coverslips are pretreated to remove RNA.

13. Add 5 µM TOTO-3 to the sample for 20 min for DNA counterstaining. 14. Add Vectashield mounting medium to the coverslip and mount on a microscope slide to help prevent photobleaching. 15. Examine samples for immunocytochemical staining. It is important to consider the intensity of staining as well as the localization of staining. SSEA-4 and the TRA antigens are all located at the cell surface, and the staining should reflect this. Conversely, Oct-4 and Nanog are both transcription factors and should be

Isolation of Embryonic Stem Cells

1A.1.7 Current Protocols in Stem Cell Biology

Supplement 1

Figure 1A.1.2 Immunocytochemical localization of the pluripotency markers (A) Oct-4 and (B) Nanog in nhpES cells. Immunocytochemistry allows for the determination of heterogeneity in colonies. (A) Oct-4 and (B) Nanog are transcription factors and should be localized to the nuclei in healthy pluripotent colonies. This staining also highlights the prominent nucleoli observed in ES cells.

localized to the nucleus (Fig. 1A.1.2). Failure to localize properly could indicate problems in the stem cell culture or the labeling protocol. SUPPORT PROTOCOL 2

DETECTION OF OCT-4, NANOG, SOX-2, AND REX-1 BY RT-PCR RT-PCR allows for the rapid screening of expression for a number of proteins in a bulk population of embryonic stem cells. This technique’s primary strength, sensitivity, is also a major limitation, as low levels of mRNA can still be amplified, resulting in a positive signal. It is also difficult to measure the expression levels across all embryonic stem cells, as high expression in one population of cells will mask low expression in another population. However it is a quick and cost-efficient means of measuring expression of pluripotent genes and is confirmatory when combined with immunocytochemistry.

Materials

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

One 70% confluent well of a 6-well plate of nhpES cells (Basic Protocol) TRIzol Reagent (Invitrogen) Chloroform (minimum 99%; Sigma) Isopropanol 75% ethanol in nuclease-free water Nuclease-free water (ISC Bioexpress; http://www.bioexpress.com) DNA-free Kit (Ambion) containing: 10× DNase I buffer recombinant DNase I (rDNase I) DNase Inactivation Reagent Reverse Transcription System (Promega) containing: 25 mM MgCl2 5× reverse transcription buffer 10 mM dNTP mixture Recombinant RNasin ribonuclease inhibitor Reverse transcriptase Oligo(dT) primer Biolase PCR Kit (Bioline) containing: Biolase Taq DNA Polymerase 10× NH4 Buffer

1A.1.8 Supplement 1

Current Protocols in Stem Cell Biology

50 mM MgCl2 Solution 2× PolyMate Additive 10 mM dNTP mix (Roche Applied Science) PCR primers for rhesus EC markers: Oct-4: forward: 5 -CGACCATCTGCCGCTTTGAG-3 reverse: 5 -CCCCCTGTCCCCCATTCCTA-3 Nanog: forward: 5 -CTGTGATTTGTGGGCCTGAA-3 reverse: 5 -TGTTTGCCTTTGGGACTGGT-3 Rex-1: forward: 5 -GCGTACGCAAATTAAAGTCCAGA-3 reverse: 5 -CAGCATCCTAAACAGCTCGCAGAAT-3 Sox2: forward: 5 -CCCCCGGCGGCAATAGCA-3 reverse: 5 -TCGGCGCCGGGGAGATACAT-3 Cell scrapers 15-ml conical tubes Refrigerated centrifuge Automatic pipettors and filtered pipet tips designated for RNA work (RNase-free; Molecular BioProducts; http://www.mbpinc.com/html/index.html 0.6-ml microcentrifuge tubes, sterile and RNase free (Molecular BioProducts; http://www.mbpinc.com/html/index.html) 0.2-ml PCR reaction tubes (ISC Bioexpress, http://www.bioexpress.com) Thermal cycler (e.g., PTC-200 Peltier Thermal Cycler; MJ Research) Additional reagents and equipment for isolating ES cells (Basic Protocol), nucleic acid quantitation (Gallagher and Desjardins, 2006) and agarose gel electrophoresis (Voytas, 2000) NOTE: Use nuclease-free water to prepare all reagents. All tubes and pipets must be RNase-free. Always wear gloves while handling samples. Do not leave tubes open any longer than absolutely necessary. Before each use, wipe gloves and pipets with RNase Away (Molecular BioProducts; http://www.mbpinc.com/html/index.html).

Isolate RNA 1. Isolate nhpES cells by manual scraping of cell colonies. Transfer cells to a 15-ml conical tube and centrifuge 5 min at 200 × g, room temperature. 2. Remove all but ∼100 µl of supernatant, add 1 ml of TRIzol to the cell pellet, and mix by vortexing for 10 sec. 3. Add 200 µl chloroform and vortex for 30 sec. 4. Centrifuge 5 min at 2500 × g, 4◦ C.

Purify RNA 5. Carefully transfer the aqueous phase (∼600 µl) to a sterile RNase-free 0.6 ml microcentrifuge tube. Avoid disturbing the white precipitate layer, which contains DNA and protein) and add 600 µl isopropanol. Incubate at −20◦ C for at least 2 hr, but preferably overnight. 6. After incubation, centrifuge tube 30 min at 14,000 to 16,000 × g, 4◦ C. 7. Carefully remove the isopropanol, leaving a small amount behind in order to avoid disturbing the pellet, if necessary. 8. Add 600 µl of 75% ethanol and centrifuge 10 min at 14,000 to 16000 × g,

4◦ C.

Isolation of Embryonic Stem Cells

1A.1.9 Current Protocols in Stem Cell Biology

Supplement 1

9. Remove the ethanol, again being careful not to disturb the pellet. Dry the pellet at room temperature (pellet should become transparent). 10. Resuspend pellet in 25 to 50 µl RNase-free water.

Treat sample with DNase 11. Add 0.1 vol of 10× DNase I buffer and 1 µl rDNase I (from the Ambion DNA-free kit) to the RNA, mix gently, and incubate at 37◦ C for 20 to 30 min. We find that we get better results if we first treat the RNA with DNase.

12. Add 2 µl or 0.1 vol (whichever is greater) of resuspended DNase Inactivation Reagent, and mix well. 13. Incubate at room temperature for 2 min, mixing occasionally. 14. Centrifuge 1.5 min at 10,000 × g, 4◦ C, and transfer the supernatant to a new sterile RNase-free tube. Determine RNA concentration by measuring A260 /A280 (Gallagher and Desjardins, 2006).

Perform reverse transcription 15. To prepare the isolated RNA for the production of cDNA, incubate 1 µg of total RNA for 10 min at 70◦ C (in thermal cycler), then microcentrifuge briefly at maximum speed and place on ice. 16. In a 0.2-ml PCR reaction tube on ice, prepare a 20-µl RT-PCR reaction by adding the following reagents in the order listed:

2.4 µl 25 mM MgCl2 4 µl 5× reverse transcription buffer 1 µl 10 mM dNTP mixture 0.5 µl recombinant RNasin ribonuclease inhibitor 1 µl reverse transcriptase 1.0 µg Oligo(dT) primer 1.0 µg total RNA (from step 15) Nuclease-free H2 O to final volume of 20 µl. 17. Incubate reaction mixture in the thermal cycler at 42◦ C for 1 hr, followed by 5 min at 95◦ C, followed immediately by 5 min at 4◦ C. The cDNA can be stored for long periods of time at −20◦ C or can be used immediately in the procedures below.

Amplify cDNA and characterize product The PCR programs described below are for an MJ Research PTC-200 Peltier Thermal Cycler. They can serve as a starting point for researchers employing other thermal cyclers. 18. Prepare a 50-µl amplification reaction by adding the following reagents (from Biolase PCR Kit, except for the 10 mM dNTP mix, which is purchased from Roche Applied Science) in the order listed:

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

5.0 µl 10× NH4 buffer 1.5 µl 50 mM MgCl2 1.0 µl 10 mM dNTP mix 0.5 µl 50 µM forward primer for marker of interest 0.5 µl 50 µM reverse primer for marker of interest 0.5 µl 5 U/µl Biolase Taq DNA polymerase 1.0 µg cDNA (from step 17) Nuclease-free H2 O to final volume of 50 µl.

1A.1.10 Supplement 1

Current Protocols in Stem Cell Biology

19a. To amplify for Oct-4 (resulting in a product that is 577 bp): Use the following thermal cycling program: 1 cycle: 35 cycles:

1 cycle

5 min 30 sec 30 sec 45 sec 5 min

94◦ C 94◦ C 60◦ C 72◦ C 72◦ C

(initial denaturation) (denaturation) (annealing) (extension) (final extension).

19b. To amplify for Nanog (resulting in a product that is 152 bp): Use the following thermal cycling program: 1 cycle: 35 cycles:

1 cycle:

5 min 30 sec 30 sec 1 min 5 min

94◦ C 94◦ C 62◦ C 72◦ C 72◦ C

(initial denaturation) (denaturation) (annealing) (extension) (final extension).

19c. To amplify for Rex-1 (resulting in a product that is 350 bp): Use the following thermal cycling program: 1 cycle: 35 cycles:

1 cycle

5 min 30 sec 30 sec 45 sec 5 min

94◦ C 94◦ C 56◦ C 72◦ C 72◦ C

(initial denaturation) (denaturation) (annealing) (extension) (final extension).

19d. To amplify for Sox-2 (resulting in a product that is 448 bp): Use the following thermal cycling program: 1 cycle: 35 cycles:

1 cycle:

5 min 30 sec 30 sec 1 min 5 min

94◦ C 94◦ C 57.9◦ C 72◦ C 72◦ C

(initial denaturation) (denaturation) (annealing) (extension) (final extension).

20. To determine presence of product and product size, load 10 µl of each product and 5 µl of a 100-bp DNA size ladder onto a 1.5% agarose gel containing 0.5 µg ethidium bromide and perform electrophoresis in 1× TAE buffer (Voytas, 2000).

KARYOTYPING OF NONHUMAN PRIMATE ES CELL CULTURES Human embryonic stem cells have well documented karyotypic instability in culture, and there is evidence suggesting that nonhuman primate ES cells have similar instability. It is therefore imperative that cultures be checked periodically (every 6 months and any time the pattern of cell growth changes). The protocol below is based on a protocol for human ES cells developed by Dr. Maya Mitalipova, Whitehead Institute for Biomedical Research, and modified for nonhuman primates in the authors’ laboratory. If the investigator does not have the interest or resources to perform this in the laboratory, samples can be sent to the University of Pittsburgh Cytogenetics Facility under the direction of Dr. Susanne Gollin (http://www.upci.upmc.edu/facilities/Cytogen/).

SUPPORT PROTOCOL 3

Isolation of Embryonic Stem Cells

1A.1.11 Current Protocols in Stem Cell Biology

Supplement 1

Materials nhpES cells cultures in log-phase growth in 6-well plates (Basic Protocol) Dulbecco’s phosphate-buffered saline (DPBS, Ca2+ - and Mg2+ -free; Invitrogen, cat. no. 14190-144) TrypLE cell dissociation enzyme (Invitrogen) nhpES cell medium (see recipe) 1 µg/ml ethidium bromide working solution (see recipe) 10 µg/ml KaryoMAX Colcemid solution (Invitrogen) Hypotonic solution: 0.075 M KCl, 37◦ C Fixative: 1:3 (v/v) acetic acid/methanol 0.025% trypsin in DPBS (prepare from 0.5% trypsin stock, see recipe) 2% (v/v) fetal bovine serum (Invitrogen, cat. no. 16000-044) in DPBS Giemsa stain solution: KaryoMAX Giemsa Stain (Invitrogen) diluted to 6% in Gurr’s buffer, pH 6.8 (see below) Gurr’s buffer, pH 6.8: dissolve one Gurr’s buffer tablet in 1 liter distilled H2 O 15-ml conical centrifuge tubes Inverted microscope Fine glass needle for dissecting ESC colonies: pull a Pasteur pipet as thinly as possible while heating over Bunsen burner, such that a pair of needles with pointed sealed ends (mandatory) are produced, and bend according to preference for optimal access to the wells; alternatively, use commercially available stem cell knives (Swemed cat. no. 25111-109M; http://www.swemed.com) Centrifuge Glass microscope slides Beaker of hot water for adjusting humidity/temperature conditions Slide warmer Coplin jars Cytovision Workstation and Genus software (Applied Imaging) or bright-field microscope with green interference filter and digital camera, with digital image processing software (e.g., Adobe Photoshop) Collect cells 1. Remove medium from three wells of a 6-well culture plate of log-phase nhpES cells. 2. Rinse wells with 37◦ C DPBS, discard, and add 1 ml of 37◦ C TrypLE to enzymatically loosen/dissociate cells (ES cells will round up in 1 to 2 min; observe with inverted microscope). Add 2 ml nhpES cell medium to inactivate TrypLE. MEFs will not dissociate during the first 1 to 2 min; therefore minimizing the time in TrypLE is important in reducing the MEF contamination in the collected ES cells.

3. Working in the original well, tease rounded-up ESC colonies into a near single-cell suspension using a fine glass needle. 4. Add sufficient 1 µg/ml ethidium bromide solution to the well (still containing the TrypLE) for a final concentration of 12 ng/ml. Incubate 40 min at 37◦ C.

Arrest mitosis 5. Add the microtubule-inhibiting compound Colcemid to this suspension to a final concentration of 120 ng/ml. Incubate 20 min at 37◦ C. Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

6. Collect the cell suspension in a 15-ml conical tube and centrifuge for 8 min at 800 × g, room temperature. 7. Remove supernatant, then add 1 ml 37◦ C DPBS to the cell pellet and centrifuge 8 min at 800 × g, room temperature.

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Current Protocols in Stem Cell Biology

8. Discard the supernatant and resuspend the pellet in 1 ml of 37◦ C TrypLE. 9. After 1 min, add 2 to 3 ml nhpES cell medium to the tube to inactivate the TrypLE and repeat centrifugation. 10. Discard supernatant and resuspend pellet thoroughly in as small a quantity of the residual supernatant as possible.

Swell the cells 11. Add 5 ml of 37◦ C hypotonic solution (0.075 M KCl). Incubate at 37◦ C for 20 min. 12. Add ∼10 drops of fixative (1:3 v/v acetic acid/methanol) to the suspension, gently invert twice to mix, and incubate 5 min at room temperature to prefix the cells. 13. Centrifuge 8 min at 800 × g, room temperature. Discard supernatant and resuspend pellet in remaining fluid.

Fix the cells 14. Add 5 ml of fixative slowly to the suspension of fragile prefixed cells while gently tapping the tube. 15. Incubate cells at room temperature for 30 min to fix, then centrifuge 8 min at 800 × g, room temperature. Discard supernatant and resuspend pellet in remaining fluid. 16. Repeat steps 14 and 15 twice more. At this point the fixed cells can be stored at −20◦ C for several weeks in fixative at ∼10,000 cells/ml before proceeding if necessary.

Prepare the slides 17. Remove supernatant from final pellet and resuspend at a concentration of ∼10,000 cells/ml in fixative. 18. Using an automatic pipettor with a 20-µl pipet tip, place 10 to 20 µl of cell suspension on a glass slide and examine at 10× magnification for quality of cell preparation, noting number of cells in mitosis and quality of chromosome spread (i.e., if chromosomes are well separated or if numerous chromosomes are lying on top of one another, hindering isolation for karyotyping). 19. Adjust the quality of the slide preparation and fine tune by adjusting humidity and/or temperature factors using a beaker of hot water and/or a slide warmer to optimize quality and spreading of chromosomes. Individual conditions will vary and investigators will need to determine the optimum conditions in their own laboratories. Further discussion of optimizing chromosome spreads may be found in Bayani and Squire (2004).

Perform GTG banding on chromosomes 20. Age prepared slides on a 75◦ C slide warmer for 1 to 2 hr, then cool to room temperature and immerse in freshly prepared 0.025% trypsin solution for 25 sec. At end of this time period, immediate immerse in 2% FBS/DPBS for 10 sec. 21. Rinse slides twice in DPBS, then immerse in Giemsa stain solution for 2 to 3 min. Rinse twice in Gurr’s buffer and finally rinse in deionized water. 22. Allow slide to air dry. 23. Analyze chromosome spreads using Applied Imaging Cytovision and Genus software according to the manufacturers instructions. Alternatively, image chromosome spreads using a 100× oil objective on a high-quality research microscope with green interference filter, and photograph, preferably using a digital camera.

Isolation of Embryonic Stem Cells

1A.1.13 Current Protocols in Stem Cell Biology

Supplement 1

Figure 1A.1.3 G-banded karyotype of a male nhpES cell line. Rhesus monkey cells have a normal karyotype of 20 autosomes and 2 sex chromosomes. The Y chromosome is particularly difficult to observe, as it is very small in this species.

Digital image processing software such as Adobe Photoshop can then isolate individual chromosomes. In this manner a simple chromosome count can be easily completed. For further analysis of correct chromosome type and number see below.

24. Arrange chromosomes in matching pairs according to accepted classifications. Chromosome designation of the rhesus macaque (Macaca mulatta; Fig. 1A.1.3) is in accordance with the Macaca mulatta chromosome classification proposed by Pearson et al. (1979). A routine mitotic cell count is 20 metaphases, analyzing chromosomes band-by-band in three cells, two to three photos, and two to three karyotypes. (ACMG, 1999). SUPPORT PROTOCOL 4

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

TERATOMA FORMATION IN NOD-SCID MICE Teratoma formation in immunocompromised mice is a classic pluripotency test and the most stringent measure of pluripotency short of contribution to chimera formation. Chimera formation is unethical using human ES cells (at least into human embryos) and not routinely practical using nhpES cells and NHP embryos. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations for the care and use of laboratory animals.

1A.1.14 Supplement 1

Current Protocols in Stem Cell Biology

Materials 5–50 × 105 exponentially growing, high-quality ES cells; typically three wells of a 6-well plate (see Basic Protocol; if possible, use cells that have been recently karyotyped; see Support Protocol 3) Normal saline (0.9% w/v NaCl), sterile Immunocompromised mice (e.g., NOD-SCID; The Jackson Laboratory), 7 weeks old Anesthetic solution: 20 mg/ml ketamine/0.5 mg/ml acepromazine in normal saline 10% (v/v) formalin (formaldehyde concentration, 3.7% v/v) in DPBS (Invitrogen, cat. no. 14190-144) 70%, 90%, 95%, and 100% ethanol Paraffin wax Hematoxylin Eosin Acid rinse: combine 500 ml distilled H2 O and 1 ml glacial acetic acid Ammonia rinse: combine 480 ml distilled H2 O and 1 ml ammonium hydroxide 1-ml syringe and 25-G needle Scalpels and scissors Peloris tissue processor (Vision BioSystems, http://www.vision-bio.com/; optional) Embedding blocks Microtome Microscope slides 1. Harvest stem cells by manual passaging (Basic Protocol), centrifuge 5 min at 200 × g, room temperature, remove supernatant, and resuspend cells in ∼400 µl of normal saline. Load stem cell suspension into 1-ml syringe and attach 25-G needle. 2. Prepare mice by i.p. injection of 100 µl anesthetic solution using a 1-ml syringe and 25-G needle. This will not completely anesthetize the mouse, but serves the purpose of relaxing the testis from the abdomen.

3. Inject 100 µl of stem cell solution into the testis of each mouse and return to cage. Alternatively cells can be injected subcutaneously in the hind quarters. On an anecdotal basis, it is believed that injection into the testis requires fewer cells for teratoma formation, but this has not been rigorously tested.

4. Monitor tumor formation daily until the tumor is palpable, typically at 12 to 16 weeks post-injection. 5. Euthanize mice by CO2 asphyxiation and dissect out tumors. 6. Place tumor in 20 ml of 10% formalin in PBS and leave for 48 to 72 hr at room temperature. Large tumors (>5 mm) should be pierced with a scalpel or scissors to allow penetration of formaldehyde into deeper tissues. Tumors should be fixed for several days to ensure adequate fixation.

7. After fixation, cut the teratomas into smaller pieces, 3 to 5 mm in diameter, and return to 10% formalin for 8 to 12 hr of further fixation. Process using a Peloris processor for dehydration and embedding or process manually as in the subsequent steps. 8. Dehydrate tissue by immersing successively for 45 min each in 70%, 90%, and 95% ethanol, then three times, each time for 45 min, in 100% ethanol. Next, immerse three time, each time for 45 min, in xylene to clear the samples, then three time, each time for 45 min in paraffin wax (melted at 56◦ to 62◦ C) to infiltrate the samples with paraffin. Finally, place samples into blocks and immerse in paraffin for sectioning. Current Protocols in Stem Cell Biology

Isolation of Embryonic Stem Cells

1A.1.15 Supplement 1

Table 1A.1.1 Hematoxylin and Eosin Staining Protocol

Step

Reagent

Time

1

Xylene

1 min

2

Xylene

1 min

3

Xylene

2 min

4

100% ethanol

30 sec

5

100% ethanol

30 sec

6

95% ethanol

25 sec

7

95% ethanol

25 sec

8

Water

20 sec

9

Hematoxylin

10 min

10

Water

10 sec

11

Water

6 min a

12

Acid rinse

13

Water

6 sec 20 sec b

14

Ammonia rinse

30 sec

15

Water

8 min

16

Eosin

3 min

17

95% ethanol

10 sec

18

95% ethanol

10 sec

19

100% ethanol

10 sec

20

100% ethanol

10 sec

21

Xylene

1 min

22

Xylene

1 min

23

Xylene

1 min

24

Xylene

1 min

a 500 ml distilled H O plus 1 ml glacial acetic acid. 2 b 480 ml distilled H O plus 1 ml ammonium hydroxide. 2

9. Cut 0.4-µm sections using a microtome and place on slides. Stain with hematoxylin and eosin using the steps and timing shown in Table 1A.1.1. It is best to collaborate with a trained pathologist/histologist to analyze the stained sections. Teratomas can be disorienting when first examined. If this is not possible the investigator should consult a reputable pathology text (Rosai, 2004)

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps unless otherwise indicated. For suppliers, see SUPPLIERS APPENDIX.

Ethidium bromide working solution, 1 µg/ml Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

Stock solution: Prepare 10 µg/ml ethidium bromide (Sigma) in Hanks’ balanced salt solution without calcium and magnesium (Invitrogen). Store up to 3 months at 4◦ C. Working solution: Add 10 ml of 10 µg/ml ethidium bromide stock solution to 90 ml sterile distilled water for a working concentration of 1 µg/ml.

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Current Protocols in Stem Cell Biology

MEF medium Dulbecco’s Modified Eagle Medium, high-glucose formulation (Invitrogen) supplemented with: 10% fetal bovine serum (FBS; Invitrogen), heat inactivated 1× Pen/Strep (add from 100× stock; Invitrogen) 1× L-glutamine (add from 100× stock; Invitrogen) 1× nonessential amino acids (add from 100× stock; Invitrogen) Filter sterilize using 0.22-µm filter Store up to 1 week at 4◦ C nhpES cell medium 80% Knockout DMEM (Invitrogen) supplemented with: 20% (v/v) Knockout Serum Replacement (Invitrogen) 1× Pen/Strep (add from 100× stock; Invitrogen) 1× L-glutamine (add from 100× stock; Invitrogen) 1× nonessential amino acids (add from 100× stock; Invitrogen) 12 ng/ml basic fibroblast growth factor (bFGF; Invitrogen) 10 ng/ml Activin A (Sigma) 10 ng/ml human leukemia inhibitory factor (hLIF; Chemicon) Filter sterilize using 0.22-µm filter Store up to 1 week at 4◦ C TALP-HEPES medium Stock solution: 114 mM NaCl 3.2 mM KCl 2 mM NaHCO3 0.4 mM NaH2 PO4 10 mM sodium lactate (add as 60% syrup) 2 mM CaCl2 0.5 mM MgCl2 10 mM HEPES 100 IU/ml penicillin 1 mg/100 ml phenol red Filter sterilize using 0.22-µm filter Store up to 1 month at 4◦ C Working solution: On day of the experiment add: 3 mg/ml BSA Fraction V (Sigma) 50 µg/ml gentamicin 60 ng/ml sodium pyruvate Filter sterilize using 0.22-µm filter Trypsin, 0.5% stock and 0.025% working solutions Stock solution (0.5% trypsin): Dilute 2.5% trypsin (Invitrogen) 1:5 in Dulbecco’s phosphate-buffered saline (DPBS; Invitrogen, cat. no. 14190-144). Store up to 6 months at −20◦ C. Working solution: (0.025% trypsin): Just before use, dilute 0.5% trypsin stock to 0.025% with DPBS. Isolation of Embryonic Stem Cells

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COMMENTARY Background Information

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

Mouse embryonic stem cells have primarily been used for the generation of improved animal models (knockouts, knockins), and in this fashion have truly transformed biomedical research. Human embryonic stem cells have the potential to similarly transform medicine by generation of cells with the potential for therapy. They also serve as a model cell for studying very early differentiation events in human embryonic and fetal development. Ethical concerns preclude the indepth examination of the pluripotency of human embryonic stem cells in chimeras, either with animal embryos or human embryos. Nonhuman primate embryonic stem cells have the potential to cross the divide between these two species and answer pluripotency questions that cannot be asked using human ES cells. If embryonic stem cells from monkeys can contribute to chimeric offspring like murine embryonic stem cells, this would allow for the development of monkey models for disease that more faithfully represent human disease. Though unlikely to completely replace mouse models due to cost and other constraints, a monkey model for aging and cognitive diseases such as Alzheimer’s would be invaluable. Monkey embryonic stem cells are also the perfect cells to use for preclinical testing of any potential therapies using human embryonic stem cells. Work on the differentiation of nhpES cells is progressing, with successful differentiation reported into neural cells (Calhoun et al., 2003; Kuo et al., 2003; Nakayama et al., 2003; Li et al., 2005), hematopoietic cells (Umeda et al., 2004, 2006), and pigmented retinal epithelium (Haruta et al., 2004). Cells differentiated into neurons have been transplanted into monkey brains (Sanchez-Pernaute et al., 2005; Takagi et al., 2005) with long-term survival, including transfer into a monkey model of Parkinson’s disease with early but promising results (Takagi et al., 2005). Monkey ES cells have been shown to contribute to chimeric embryos (Takada et al., 2002; Mitalipov et al., 2006) but no contribution has been shown in fetuses or offspring to date. It is well known in the mouse embryonic stem cell field that ES cells can maintain pluripotent markers but fail to contribute to chimeric tissues or the germ line. Therefore, it may be necessary to screen dozens of nhpES

cell lines before one is found capable of this task. The derivation of nonhuman primate ES cells has continued successfully but sporadically since the first isolation (Thomson et al., 1995). nhpES cells have been isolated from in vivo–derived embryos (Thomson et al., 1995, 1996) and in vitro embryos including those derived by intracytoplasmic sperm injection (ICSI; Suemori et al., 2001; Mitalipov et al., 2006; Navara et al., 2007). They have even been derived from parthenogenetic embryos (Cibelli et al., 2002). Derivations include three different nonhuman primate species, rhesus monkey (Macaca mulatta; Thomson et al., 1995; Mitalipov et al., 2006; Navara et al., 2007), cynomolgus monkey (Macaca fascicularis; Suemori et al., 2001; Cibelli et al., 2002), and marmoset (Callithrix jacchus; Thomson et al., 1996; Sasaki et al., 2005).

Critical Parameters and Troubleshooting Before attempting to isolate nhpES cells, investigators should develop the techniques for passaging existing human or monkey embryonic stem cells. Many of the steps require an understanding of the pluripotent phenotype for selection of the highest-quality cells. It would be unfortunate to incur the time and expense of generating NHP embryos and attempting to isolate stem cells, only to lose them as a result of failure to recognize the cells in culture or errors in passaging or preparing mouse embryonic feeder cells. The nhpES cell medium described in this unit (see recipe) was developed based on published reports that Activin A (Vallier et al., 2005) and increased levels of bFGF (Xu et al., 2005; Levenstein et al., 2006) are helpful in maintaining pluripotency. Additionally, although leukemia inhibitory factor has been shown to be extraneous for pluripotency, most derivation media include this component. The nhpES medium described in this unit has been successfully used in the authors’ laboratory, but it is rather costly. Other derivation media have been described (Thomson et al., 1995; Suemori et al., 2001; Sasaki et al., 2005; Mitalipov et al., 2006) for nhpES cells, and investigators may want to look into these if costs warrant. Embryo quality most likely plays a large role in the success of stem cell derivation. In the authors’ research, it has been found that

1A.1.18 Supplement 1

Current Protocols in Stem Cell Biology

the embryos that develop fastest in vitro also yield the highest success rate for embryonic stem cell derivation (Navara et al., 2007). This correlates with a long-standing belief by reproductive biologists that the highest-quality embryos also develop the fastest. The authors have isolated stem cells from later-developing embryos, but at rates one-half of that obtained with the more rapidly developing embryos. While it has been shown to be possible to isolate ES cell lines even from embryos believed to have arrested (Zhang et al., 2006), beginning investigators will want to ensure that they are starting with only the best embryos. The authors retain all early cultures, even those from which passaging has been performed, for an additional 2 weeks after passaging to ensure that all potential stem cells have been harvested. Once cell lines are established, they should be frozen early and often. As soon as the cells exist in multiple cultures, they should be cryopreserved. This is a necessary step for safeguarding against contamination, aneuploidy (see below), or other culture errors. Perhaps the biggest risk in the culture of embryonic stem cells, particularly for investigators just beginning to culture these cells, is the risk of cells becoming aneuploid in culture. Embryonic stem cells should be tested every 6 months for proper and stable karyotype, and should also be checked when growth conditions change, e.g., in cases where there is faster growth or less differentiation than expected. In order to ensure the highest-quality immunocytochemistry, cells should be fixed in 37◦ C formaldehyde as soon as possible after removal from the incubator (within 1 or 2 min). It is best to process the staining all at once, instead of stopping at any given step, and slides should be examined as soon after staining as possible. Commercial antibodies may change over time such that the antibody purchased 6 months ago is not the same antibody purchased today. This can be due to a change in the lot of antibody or a complete reworking of the antibody from the vendor. If an antibody stops working, it will be necessary to test various fixations and antibody dilutions to reoptimize the labeling conditions. Karyotyping of any cell type requires some adjustments to the system, and this is especially true of embryonic stem cells. Several factors can reduce optimal chromosome spreading and banding, and this in turn can inhibit proper interpretation. If not enough mitotic figures are observed, the concentration and incubation time of Colcemid treatment

can be increased. Conversely, if very short chromosomes result, this is generally a sign of too much Colcemid or too long an incubation. Chromosomes can also be lengthened by increasing the ethidium bromide concentration or incubation time. Fine tuning the slide preparation conditions by modifying the humidity or temperature or by varying the exposure time to hypotonic solution can increase the quality of chromosome spreads. Poor banding is usually a result of over- or under-trypsinization. When adjusting the conditions, trypsin exposure time should be varied by 2-sec intervals. Bands that are not distinct, with diffuse chromosomes, mean that trypsin time should be decreased; conversely, metaphase chromosomes with few light bands indicate that increased time with trypsin is needed. When interpreting the karyotype, random chromosome loss should not be a concern unless three cells are detected with the same hypodiploidy. If a single hyperploid or aneuploid cell is observed, 20 more cells should be counted. If another identical karyotype is found, it is likely a clone. A repeat karyotype should be performed on the cells to monitor clonal propagation in culture. Aneuploid cells very often have exaggerated pluripotency characteristics, and are thus likely to be selected by manual passaging, making it possible for them to quickly overrun the colony. If this happens, return to an earlier passage from the freezer and throw out the cultures displaying aneuploidy. Alternatively, if no earlier passages exist, single cells can be isolated using a cell sorter, and clones grown from these single cells can be analyzed for pluripotency and proper karyotype. This procedure is incredibly inefficient, but could be used to save a precious cell line. If teratomas fail to form, the number of cells injected can be increased. This may be an effect of viability after harvesting, and this can be tested using a simple live/dead stain such as trypan blue. Cells for teratoma formation should be of the same high quality as those used for other pluripotency assays. Resist the temptation to use already differentiating cells with the justification that they are going to differentiate anyway.

Anticipated Results Investigators with a successful history maintaining or propagating existing human or nonhuman primate ES cells should be able to successfully isolate embryonic stem cell lines from 25% to 50% of fully expanded

Isolation of Embryonic Stem Cells

1A.1.19 Current Protocols in Stem Cell Biology

Supplement 1

blastocysts. Faithful attention to manual passaging of only the highest-quality cells should allow for greater than 75% of early established lines to be propagated to stability. Cells with the proper morphology (closely packed cells with a high nucleus:cytoplasm ratio and prominent nucleoli) will display most, if not all, of the described markers for pluripotency and will acquire the others in culture.

Time Considerations

Derivation and Characterization of Non-Human Primate Embryonic Stem Cells

The process of immunosurgery requires ∼2 hr. At a time point 48 hr prior to the day of immunosurgery, 6-well plates containing feeder cells should be prepared, and 2 hr before the immunosurgery, the feeder cell medium should be replaced with nhpES cell medium. Attachment of the isolated ICM takes between 24 and 72 hr. Investigators can wait longer, but the success rate of derivation of an ES line from embryos that take longer than 72 hr to attach approaches zero. It is ∼2 weeks from the time of immunosurgery until the derived ES cells are ready for passaging. After this point, they should be passaged every 5 to 7 days. Immunocytochemical staining takes ∼4 hr, not including the overnight incubation for Oct4 and Nanog antibodies. RT-PCR analysis of pluripotency can be completed in 6 to 8 hr on a single day, or can be split overnight so that the first day includes RNA isolation, requiring about 45 min, and the next day requires 2 to 3 hr for generating cDNA, performing PCR, and analyzing by gel electrophoresis. Karyotyping requires 6 to 8 hr on the first day for harvesting the ES cells, fixing them, and preparing glass slides. The next two steps can be completed in 1 day or split over 2 days for convenience. G-banding of the prepared slides requires ∼4 hr; allow at least another 4 hr for analysis of the prepared slides, depending on how many slides have been prepared and the familiarity of laboratory personnel with cytogenetic analysis. Preparing the cells for teratoma formation requires ∼1 hr, and injection into an immunocompromised mouse requires another hour. Teratomas require at least 8 weeks to develop, and generally require more than 12 weeks to develop in vitro. Investigators should not try to speed this process by injecting a larger number of cells. The teratoma will become large more quickly but the individual cell types will not have enough time to differentiate; it is difficult to interpret poorly differentiated teratomas.

Literature Cited American College of Medical Genetics (ACMG). 1999. Standards and Guidelines for Clinical Genetics Laboratories. 2nd ed. ACMG, Rockville, Md. Bayani, J. and Squire, J.A. 2004. Preparation of cytogenetic specimens from tissue samples. Curr. Protoc. Cell. Biol. 23:22.2.1-22.2.15. Calhoun, J.D., Lambert, N.A., Mitalipova, M.M., Noggle, S.A., Lyons, I., Condie, B.G., and Stice, S.L. 2003. Differentiation of rhesus embryonic stem cells to neural progenitors and neurons. Biochem. Biophys. Res. Commun. 306:191-197. Cibelli, J.B., Grant, K.A., Chapman, K.B., Cunniff, K., Worst, T., Green, H.L., Walker, S.J., Gutin, P.H., Vilner, L., Tabar, V., Dominko, T., Kane, J., Wettstein, P.J., Lanza, R.P., Studer, L., Vrana, K.E., and West, M.D. 2002. Parthenogenetic stem cells in nonhuman primates. Science 295:819. Gallagher, S.R. and Desjardins, P.R. 2006. Quantitation of DNA and RNA with absorption and fluorescence spectroscopy. Curr. Protoc. Mol. Biol. 76:A.3D.1-A.3D.21. Haruta, M., Sasai, Y., Kawasaki, H., Amemiya, K., Ooto, S., Kitada, M., Suemori, H., Nakatsuji, N., Ide, C., Honda, Y., and Takahashi, M. 2004. In vitro and in vivo characterization of pigment epithelial cells differentiated from primate embryonic stem cells. Invest. Ophthalmol. Vis. Sci. 45:1020-1025. Hewitson, L. 2004. Primate models for assisted reproductive technologies. Reproduction 128:293-299. Kuo, H.C., Pau, K.Y., Yeoman, R.R., Mitalipov, S.M., Okano, H., and Wolf, D.P. 2003. Differentiation of monkey embryonic stem cells into neural lineages. Biol. Reprod. 68:1727-1735. Levenstein, M.E., Ludwig, T.E., Xu, R.H., Llanas, R.A., VanDenHeuvel-Kramer, K., Manning, D., and Thomson, J.A. 2006. Basic fibroblast growth factor support of human embryonic stem cell self-renewal. Stem Cells 24:568-574. Li, T., Zheng, J., Xie, Y., Wang, S., Zhang, X., Li, J., Jin, L., Ma, Y., Wolf, D.P., Zhou, Q., and Ji, W. 2005. Transplantable neural progenitor populations derived from rhesus monkey embryonic stem cells. Stem Cells 23:1295-1303. Maatman, R., Gertsenstein, M., de Meijer, E., Nagy, A., and Vintersten, K. 2003. Aggregation of embryos and embryonic stem cells. Methods Mol. Biol. 209:201-230. Mitalipov, S., Kuo, H.C., Byrne, J., Clepper, L., Meisner, L., Johnson, J., Zeier, R., and Wolf, D. 2006. Isolation and characterization of novel rhesus monkey embryonic stem cell lines. Stem Cells 24:2177-2186. Nakayama, T., Momoki-Soga, T., and Inoue, N. 2003. Astrocyte-derived factors instruct differentiation of embryonic stem cells into neurons. Neurosci. Res. 46:241-249. Navara, C.S., Mich-Basso, J., Redinger, C., Ben-Yehudah, A., and Schatten, G. 2007. Pedigreed non-human primates embryonic stem cells

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display family and sex related differences in gene expression. Submitted for publication.

onic stem cell line. Proc. Natl. Acad. Sci. U.S.A. 92:7844-7848.

Pearson, P.L., Roderick, T.M., Davisson, M.T., Garver, J.J., Warburton, D., Lalley, P.A., and O’Brien, S.J. 1979. Report of the committee on comparative mapping. Cytogenet. Cell Genet. 25:82-95.

Thomson, J.A., Kalishman, J., Golos, T.G., Durning, T.G., Harris, C.P., and Hearn, J.P. 1996. Pluripotent cell lines derived from common marmoset (Callithrix jacchus) blastocysts. Biol. Reprod. 55:254-259.

Rosai, J (ed.). 2004. Rosai and Ackerman’s Surgical Pathology. 9th ed. Elsevier, New York.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

Sanchez-Pernaute, R., Studer, L., Ferrari, D., Perrier, A., Lee, H., Vinuela, A., and Isacson, O. 2005. Long-term survival of dopamine neurons derived from parthenogenetic primate embryonic stem cells (cyno-1) after transplantation. Stem Cells 23:914-922. Sasaki, E., Hanazawa, K., Kurita, R., Akatsuka, A., Yoshizaki, T., Ishii, H., Tanioka, Y., Ohnishi, Y., Suemizu, H., Sugawara, A., Tamaoki, N., Izawa, K., Nakazaki, Y., Hamada, H., Suemori, H., Asano, S., Nakatsuji, N., Okano, H., and Tani, K. 2005. Establishment of novel embryonic stem cell lines derived from the common marmoset (Callithrix jacchus). Stem Cells 23:13041313. Schatten, G., Smith, J., Navara, C., Park, J.H., and Pedersen, R. 2005. Culture of human embryonic stem cells. Nat. Methods. 2:455-463. Suemori, H., Tada, T., Torii, R., Hosoi, Y., Kobayashi, K., Imahie, H., Kondo, Y., Iritani, A., and Nakatsuji, N. 2001. Establishment of embryonic stem cell lines from cynomolgus monkey blastocysts produced by IVF or ICSI. Dev. Dyn. 222:273-279. Takada, T., Suzuki, Y., Kondo, Y., Kadota, N., Kobayashi, K., Nito, S., Kimura, H., and Torii, R. 2002. Monkey embryonic stem cell lines expressing green fluorescent protein. Cell Transplant 11:631-635. Takagi, Y., Takahashi, J., Saiki, H., Morizane, A., Hayashi, T., Kishi, Y., Fukuda, H., Okamoto, Y., Koyanagi, M., Ideguchi, M., Hayashi, H., Imazato, T., Kawasaki, H., Suemori, H., Omachi, S., Iida, H., Itoh, N., Nakatsuji, N., Sasai, Y., and Hashimoto, N. 2005. Dopaminergic neurons generated from monkey embryonic stem cells function in a Parkinson primate model. J. Clin. Invest. 115:102-109. Thomson, J.A., Kalishman, J., Golos, T.G., Durning, M., Harris, C.P., Becker, R.A., and Hearn, J.P. 1995. Isolation of a primate embry-

Umeda, K., Heike, T., Yoshimoto, M., Shiota, M., Suemori, H., Luo, H.Y., Chui, D.H., Torii, R., Shibuya, M., Nakatsuji, N., and Nakahata, T. 2004. Development of primitive and definitive hematopoiesis from nonhuman primate embryonic stem cells in vitro. Development 131:18691879. Umeda, K., Heike, T., Yoshimoto, M., Shinoda, G., Shiota, M., Suemori, H., Luo, H.Y., Chui, D.H., Torii, R., Shibuya, M., Nakatsuji, N., and Nakahata, T. 2006. Identification and characterization of hemoangiogenic progenitors during cynomolgus monkey embryonic stem cell differentiation. Stem Cells 24:1348-1358. Vallier, L., Alexander, M., and Pedersen, R.A. 2005. Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell. Sci. 118:44954509. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Vrana, K.E., Hipp, J.D., Goss, A.M., McCool, B.A., Riddle, D.R., Walker, S.J., Wettstein, P.J., Studer, L.P., Tabar, V., Cunniff, K., Chapman, K., Vilner, L., West, M.D., Grant, K.A., and Cibelli, J.B. 2003. Nonhuman primate parthenogenetic stem cells. Proc. Natl. Acad. Sci. U.S.A. 100:11911-11916. Xu, R.H., Peck, R.M., Li, D.S., Feng, X., Ludwig, T., and Thomson, J.A. 2005. Basic FGF and suppression of BMP signaling sustain undifferentiated proliferation of human ES cells. Nat. Methods. 2:185-190. Zhang, X., Stojkovic, P., Przyborski, S., Cooke, M., Armstrong, L., Lako, M., and Stojkovic, M. 2006. Derivation of human embryonic stem cells from developing and arrested embryos. Stem Cells 24:2669-2676.

Isolation of Embryonic Stem Cells

1A.1.21 Current Protocols in Stem Cell Biology

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Derivation of hESC from Intact Blastocysts

UNIT 1A.2

Dusko Ilic,2 Olga Genbacev,2 and Ana Krtolica1, 3 1

StemLifeLine Inc., San Carlos, California University of California, San Francisco, California 3 Lawrence Berkeley National Laboratory, Berkeley, California 2

ABSTRACT This unit describes protocols for culturing human embryos and deriving human embryonic stem cells from the intact blastocyst. Description of the culturing begins with methods for obtaining human blastocysts using pronuclear or cleavage stage embryos left over after in vitro fertilization. Then there is a description of methods that can be used to derive human embryonic stem cell lines from the blastocyst without trophectoderm removal. Also included is a discussion of the critical steps and parameters such as zona pellucida removal, embryo quality assessment, feeder selection, when and how to transfer early embryonic outgrowths. In addition, there are protocols for embryo thawing, seeding of feeder cells, gelatin coating of plates, cleavage and blastocyst stage embryo grading, preparation and storage of reagents and solutions. Finally, there is a discussion of alternative derivation approaches as well as the timeline for the procedures. Curr. C 2007 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 1:1A.2.1-1A.2.18.
Keywords: human embryonic stem cells (hESC) r inner cell mass (ICM) r trophectoderm (TE) r zona pellucida removal r feeders

INTRODUCTION Like mouse embryonic stem cells, human embryonic stem cells (hESC) are derived from the inner cell mass (ICM) of pre-implantation embryos and can give rise to cells from all three germ layers (pluripotency). If properly maintained, they can be grown in culture virtually indefinitely while retaining their pluripotency and unlimited self-renewal capacity. It is these characteristics that make hESC ideal candidates for drug testing and future cell replacement therapies. Because hESC share these characteristics with the early embryo cells from which they originate, they can also serve as good models for studies of early human development. This is an understudied area of research because of the limited availability of the relevant tissue material as well as a variety of ethical issues related to its use. This unit describes protocols related to the derivation of pluripotent embryonic stem cells from human embryos left over after in vitro fertilization (IVF). The Basic Protocol describes a method for deriving embryonic stem cells from the intact zona pellucida–free blastocyst. The authors have used this method (previously described in Genbacev et al., 2005) to derive more than ten hESC lines. Support Protocol 1 describes culturing of embryos from either pronuclear (day 1 single-cell) or cleavage (day 3 8-cell) stage to blastocyst stage followed by zona pellucida removal by acid hydrolysis using Tyrode’s solution (Support Protocol 2) or by enzymatic digestion with pronase (Support Protocol 3). In addition, protocols are provided for embryo thawing (Support Protocol 4) and seeding of feeder cells (Support Protocol 5). NOTE: All procedures described in this unit, including preparation of reagents and solutions, should be performed under sterile culture conditions in either Class II biological

Current Protocols in Stem Cell Biology 1A.2.1-1A.2.18 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01a02s1 C 2007 John Wiley & Sons, Inc. Copyright


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1A.2.1 Supplement 1

safety cabinets or laminar flow hoods. For handling embryos, a dissecting microscope should be placed within a laminar flow hood, and a face mask should be worn to prevent contamination. NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: The described protocols usually require ethics approval from the appropriate institutional review board or equivalent entity. Typically, only embryos donated for research with consent from IVF patients can be used. Regulations may vary depending on geographic area, so inquire locally before initiating this type of research. BASIC PROTOCOL

HUMAN EMBRYONIC STEM CELL (hESC) DERIVATION Zona pellucida–free blastocysts are cultured on feeder layers in the presence of human recombinant basic fibroblast growth factor (bFGF) to allow the outgrowth of hESCs.

Materials KSR embryo culture medium supplemented with 25 ng/ml bFGF (see recipe) Zona pellucida–free blastocyst-stage embryos (Support Protocols 2 and 3) 26-G needle, sterile The Stripper micropipettor (MidAtlantic Diagnostics MXL3-STR) and 600-µm polycarbonate tips (MidAtlantic Diagnostics MXL3-600) 1.8-ml cryovials Additional reagents and equipment for preparing feeder cells in 4- or 6-well tissue culture plates (Support Protocol 5) Prepare culture plates 1. Prepare feeder cells in 4-well tissue culture plates (Support Protocol 5) 1 to 3 days before plating the zona pellucida–free blastocyst-stage embryo. Alternatively, 6-well plates may be used. Production of feeder plates should be scheduled to provide freshly plated feeder cells for transfers (see step 7). It is always better to have more wells with freshly plated feeder cells than required for embryos and transfers; plating may sometimes yield wells where feeder cells are not uniformly distributed, and these wells should not be used.

2. One to twelve hours before plating blastocysts, replace the fibroblast medium with KSR embryo culture medium supplemented with 25 ng/ml bFGF (0.5 ml/well for 4-well plates and 3.5 ml/well for 6-well plates).

Establish inner cell mass growth 3. Place the zona pellucida–free blastocyst-stage embryos in the wells of the 4-well plates prepared in step 2 (one embryo per well) and incubate at 37◦ C in 5% CO2 . Because each embryo has different genetic material, each must be plated in a separate well. The zona pellucida–free blastocyst-stage embryo should attach to feeder cell layer within 48 hr after plating (Fig. 1A.2.1). Derivation of hESCs from Intact Blastocysts

4. Replace the KSR embryo culture medium supplemented with 25 ng/ml bFGF every second day. Observe for growth up to 1 month.

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Current Protocols in Stem Cell Biology

Figure 1A.2.1 layer (f).

Zona pellucida–free blastocyst-stage embryo (e) attached to the feeder cell

Trophectoderm cells will form the first outgrowth. Extensive secretion from trophectoderm outgrowth sometimes denudes the area of feeder cells. In such cases, trophectoderm outgrowth should be disaggregated with a sterile needle, usually 6 or 7 days after plating (Fig. 1A.2.2). Because the medium does not support the growth of trophectoderm, it dies off within 10 to 14 days after plating of the embryo.

5. Once ICM outgrowth is observed (∼15 to 24 days; Fig. 1A.2.3), replace the medium and dissect the outgrowth into smaller pieces using a sterile needle. Movement of the medium in the well while transferring the dish back into the incubator separates the dissected pieces and moves them away from the original outgrowth. ICM outgrowth is usually distinguishable 15 to 24 days after plating zona pellucida–free blastocysts on feeder cell layer. At that time, the initial trophectoderm outgrowth will die off. Although by definition feeder cells should not be able to proliferate, in some cases a few cells might escape mitotic inactivation with mitomycin C or irradiation (see Support Protocol 5) and can proliferate and fill the well with feeder cells after prolonged culture. Growth of feeder cells will quickly deplete culture medium of the growth factors and nutrients; if feeder cells continue to grow, the medium should be replaced on a daily basis. However, if the growth of feeder cells is prominent, a higher dose of irradiation or mitomycin C should be used for their mitotic inactivation (see Support Protocol 5).

6. Continue to replace KSR embryo culture medium supplemented with 25 ng/ml bFGF every second day and check for growth. Dissect outgrowth again, if present. Leave the clumps in the same well until feeders start detaching from the edges of the well or the well is filled with colonies (see Fig. 1A.2.4). The time it takes to reach the point where the hESC are expanded into new wells depends on how fast the hESC divide; all hESC do not proliferate at the same rate. The timing for transfer and/or expansion of colonies varies. For example, if there is one slowly growing colony in one well, when the colony is large enough to be dissected it would be best to transfer pieces of it into a new well with fresh feeders. If a colony is still small and feeders start to deteriorate, the colony is transferred to new feeders without

Isolation of Embryonic Stem Cells

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Figure 1A.2.2 Dissection of trophectoderm outgrowth from the attached embryo. (A) Initial trophectoderm outgrowth (arrowheads). Arrow points to areas denuded of feeder cell layer (f) due to proteolytic activity of trophectoderm cells. (B) Disaggregation of the initial trophectoderm outgrowth with a needle (n). (C) Appearance of the area after dissection.

splitting. On the other hand, if there is one fast-growing colony in one well, the colony might be dissected once or twice and the pieces left in the same well until the feeders start to deteriorate. The viability of the feeder cells can also determine when the hESC colonies are transferred. Detachment of the feeder cells indicates that the cells have aged and that their value as growth-supporting cells has decreased. Some feeders can support hESC for 4 weeks before they deteriorate; others last only 2 weeks.

7. At that time transfer the colonies from each well of the 4-well tissue culture plate into a feeder-containing well of a 6-well tissue culture plate. Derivation of hESCs from Intact Blastocysts

The larger surface area in the 6-well plate allows growth of more colonies. Alternatively, 4-well tissue culture plates can also be used for propagating colonies.

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Figure 1A.2.3 Initial ICM outgrowth (io). Visible are the denuded area due to extensive proteolytic secretion of trophectoderm cells (d), feeder cell layer (f), needle scratches remaining from disaggregation of the trophectoderm (s), and dead trophectoderm cell (t).

Figure 1A.2.4

Embryonic stem cell colonies (c) on feeder cell layer (f).

During transfer of hESC colonies from one well to another, adjacent feeder cells will be transferred, too. Because there is only small number of such cells and they do not proliferate they will not interfere with further growth and culture of hESC colonies. Never combine colonies from different embryos in one well because each embryo has its own unique genetic material.

8. Repeat dissection of the colonies until there are at least two wells of the 6-well tissue culture plate with 20 colonies per well. The hESC colonies should be propagated as described (also see UNIT 1C.1) until their number is sufficient for freezing (20 to 50 colonies/cryovial).

9. Place cells from at least one well of the 6-well tissue culture plate into one cryovial (minimum of 20 colonies per vial) for freezing (see Phelan, 2006). 10. Continue to expand cells from the other wells for additional frozen cultures and for quality control. Do not discard the well from which the original colonies were dissected for at least a week because new colonies may emerge. Whenever possible, dissect only a part of the colony leaving the other part intact, until a sufficient number of wells with colonies is established (three to four wells of the 6-well plate). When large areas in the wells lack feeder cells or feeder cells look unhealthy and start detaching and dying, dispose of the plate. Usually, if feeder cells are of good quality, they

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can serve their purpose up to 1 month. It is strongly recommended that fresh feeder cells are always plated for each passage of hESC into a new well. A number of parameters can be evaluated for quality control, depending on the investigator and research being performed, e.g., morphology, proliferation rate, expression and localization of hESC markers, karyotype, telomerase activity, and the ability to differentiate into three germ layers. SUPPORT PROTOCOL 1

IN VITRO DEVELOPMENT OF BLASTOCYSTS The blastocyst is the first stage of the human embryo at which two unquestionably distinct cell populations exist: an outer cell layer or trophectoderm and a compact inner cell population called the inner cell mass (ICM). Outgrowth of the ICM cells in culture gives rise to embryonic stem cells. During the cleavage and morula stages of embryo development, differentiation into trophectoderm and ICM is still uncertain. Culturing to the blastocyst stage helps eliminate developmentally arrested embryos and increases chances for successful hESC derivation.

Materials Appropriate cell culture medium: G-1 v3 Plus medium (Vitrolife) for the 1- to 8-cell stage (day 1 pronuclear to day 3 cleavage); G-2 v3 Plus blastocyst medium (Vitrolife) for the 8-cell (day 3 cleavage) to blastocyst (day 5 or 6) stage Oil for embryo culture (sterile light mineral oil; Irvine Scientific) Pronuclear or cleaving embryos from IVF, fresh or frozen (see Support Protocol 4 for thawing directions) 6-cm tissue culture–treated plastic dish (e.g., Falcon 3046) The Stripper micropipettor (MidAtlantic Diagnostics MXL3-STR) with 135-µm and 600-µm polycarbonate tips (MidAtlantic Diagnostics MXL3-135 and MXL3-600) 1. Place six to seven 30- to 35-µl droplets of the appropriate cell culture medium in a 6-cm tissue culture dish and cover with 5 ml of oil for embryo culture (Fig. 1A.2.5). The number of droplets depends on how many embryos will be thawed. To be on the safe side, it is always good to place more drops than necessary. Oil for embryo culture is a sterile light mineral oil and is intended for use as an overlay when culturing cells in reduced volumes of medium to prevent evaporation and insulate the medium from changes in osmolarity and pH.

2. Equilibrate medium droplets by preincubating 1 to 3 hr at 37◦ C in a 5% CO2 incubator. 3. Attach a 135-µm tip to The Stripper micropipettor (Fig. 1A.2.6) and moisten with cell culture medium as follows:

Derivation of hESCs from Intact Blastocysts

a. Carefully attach a sterile Stripper tip to the stainless steel plunger by loosening the knurled collet and depressing the finger pad until the plunger protrudes 0.5 to 1.0 cm past the collet. b. Slip on the new tip and push it firmly along the plunger until it stops against the O rings at the tip of the barrel. c. Tighten the collet. d. Rinse the tip by depressing the plunger until the finger pad contacts the spring housing; immerse the tip into a drop of medium, and slowly release the plunger. Expel the medium by depressing the plunger as before. e. To expel any residual medium in the tip, push the finger pad until it enters the spring housing.

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Figure 1A.2.5 Schematic drawing of a dish containing drops of embryo culture medium covered with oil. Label each drop clearly on bottom of the dish.

Figure 1A.2.6

The Stripper micropipettor with tip used for manipulating embryonic cells.

f. Repeat this process a few times to ensure the polycarbonate tip is sufficiently moistened. The Stripper micropipettor is a precision instrument designed to manipulate gametes or embryos with a minimal amount of fluid transfer. Once the tip has been rinsed, the embryos can be manipulated. Make sure that the bore of the tip is appropriate for the diameter of the embryo by placing the tip next to the embryo and ascertaining that the inner diameter of the tip will not cause major distortion of the embryo as it is pipetted in and out of the tip. Practice, using discarded mouse, bovine, or hamster eggs/embryos, is recommended.

4. Using the moistened pipettor tip, transfer one to four embryos from the same donor into each drop of the 37◦ C equilibrated embryo culture medium under oil. Both fresh and frozen embryos can be used to obtain blastocysts. For thawing frozen embryos see Support Protocol 4.

5. Examine each embryo under the microscope (100×) and assign a grade (see Fig. 1A.2.7). The embryos with better grades (1 or 2) are more likely to develop into blastocysts. Also, low oxygen tension (5% O2 ) and low illumination (20 lux from the ceiling and 100 lux from the microscope) throughout embryo manipulation may improve the blastulation rate (Noda et al., 1994). Special low-oxygen cell incubators are available from various manufacturers.

6. Place the dish at 37◦ C in 5% CO2 and transfer embryos every 24 to 36 hr into fresh droplets of the embryo culture medium under oil (prepared as described in steps 1 and 2). When embryos start to expand in size, transfer them with a 600-µm tip instead of the 135-µm tip. 7. When the embryos reach the blastocyst stage, proceed with zona pellucida removal (Support Protocol 2 or 3). Isolation of Embryonic Stem Cells

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Figure 1A.2.7 Grading criteria for embryos at the cleavage stage (day 3 embryos). Grade 1: Equal size blastomeres without any cell fragmentation. Grade 2: Equal size blastomeres with some cell fragmentation. Grade 3: Unequal size blastomeres with no or little cell fragmentation or equal size blastomeres with moderate cell fragmentation. Grade 4. Unequal size blastomeres with moderate fragmentation or massive cell fragmentation regardless of blastomere size. Gray shading indicates nonviable cells.

SUPPORT PROTOCOL 2

REMOVAL OF THE ZONA PELLUCIDA WITH ACIDIFIED TYRODE’S SOLUTION The zona pellucida is a protective extracellular glycoprotein matrix layer surrounding oocytes and pre-implantation embryos. As the embryo grows, the zona pellucida becomes thinner, and prior to implantation into the uterine wall, the embryo hatches out of the zona pellucida completely. Assisted hatching (in vitro removal of zona pellucida) can be accomplished in several different ways. This protocol describes removal of the zona pellucida with acidified Tyrode’s solution. Removal using pronase treatment is detailed in Support Protocol 3.

Materials KSR embryo culture medium with and without 25 ng/ml bFGF (see recipe) Acidified Tyrode’s solution (Irvine Scientific) Embryos in culture (Support Protocol 1) 4-well tissue culture plate with feeder cells (Support Protocol 5) G-2 v3 Plus blastocyst medium (Vitrolife) 6-cm tissue culture dish with cell culture–treated surface (e.g., Falcon 3046) The Stripper micropipettor (MidAtlantic Diagnostics MXL3-STR) with 600-µm (MidAtlantic Diagnostics MXL3-600) and other appropriate size tips Microscope with camera 1. Place six separate 50-µl drops of KSR embryo culture medium on a cell culture– treated surface of a sterile 6-cm tissue culture dish. 2. Place two 50-µl drops of acidified Tyrode’s solution in the same dish; mark the drops of acidified Tyrode’s solution to avoid error. CAUTION: Acidified Tyrode’s solution has a pH of 2.1 to 2.5. Use appropriate precautions in handling it. One dish with the drops of KSR medium and Tyrode’s solution should be prepared for each embryo to be treated. Derivation of hESCs from Intact Blastocysts

3. Remove the embryo from the culture drop under oil using The Stripper micropipettor with an appropriate size tip and transfer it into a drop of KSR embryo culture medium.

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Current Protocols in Stem Cell Biology

Figure 1A.2.8 Grading examples for embryos at the blastocyst stage. Blastocyst-stage embryo score is a number based on the morphology, size of the inner cell mass (i), and the viability of cells as judged under the microscope on the indicated days after in vitro fertilization, according to the following rules: 1 = fully expanded or hatching on day 5; 2 = fully expanded or hatching on day 6 or moderate expansion on day 5; 3 = moderate expansion on day 6 or early cavitation on day 5; 4 = early cavitation day 6 or morula on day 5 or 6. Add to the number score (1 to 4) two alphabetic scores: the first one to grade inner cell mass (i) and the second one to grade trophectoderm (t) according to the following rules: A = large inner cell mass or continuous trophectoderm with good cell-cell adhesion; B = medium inner cell mass or areas in trophectoderm with poor cell-cell adhesion; C = no visible inner cell mass or sparse granular trophectoderm cells. Featured examples: 2AA, fully expanded blastocyst on day 6 with a large inner cell mass and continuous trophectoderm; 3AB, moderately expanded blastocyst on day 6 with a large inner cell mass and discontinuous trophectoderm; 3BB, moderately expanded blastocyst on day 6 with a poor inner cell mass and discontinuous trophectoderm; 4CC, moderately expanded blastocyst on day 6 with no visible inner cell mass or distinguishable trophectoderm. z, zona pellucida.

4. Examine the blastocyst-stage embryo under the microscope, record an image, and assign a grade (see Fig. 1A.2.8). Do not treat embryos that have initiated hatching (Fig. 1A.2.9) with acidified Tyrode’s solution. Instead, transfer them onto feeders in G-2 v3 Plus medium and place in the cell incubator. Replace the G-2 v3 Plus medium with KSR embryo culture medium supplemented with 25 ng/ml bFGF once the embryo has completely hatched and detached from the zona pellucida (from 2 to 12 hr).

5. Transfer the embryo into the first drop of acidified Tyrode’s solution for a brief rinse, and then transfer to the second drop of acidified Tyrode’s solution. Watch carefully for the dissolution of the zona pellucida (5 to 30 sec). 6. As soon as the zona pellucida is dissolved, quickly rinse the embryo by pipetting it up and down in the first drop of KSR embryo culture medium using The Stripper micropipettor with a 600-µm tip. 7. Transfer the embryo into the next drop and repeat the procedure until embryo reaches the sixth drop. Examine the embryo to ensure that the zona pellucida was completely removed (Fig. 1A.2.10). 8. Place the zona pellucida–free embryo into a well of 4-well tissue culture plate with feeder cells in 0.5 ml KSR embryo culture medium supplemented with 25 ng/ml bFGF. Place only one embryo into each well. Because each embryo has its own unique genetic material, it is crucial not to mix them.

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Figure 1A.2.9 Hatching blastocyst-stage embryo. z, zona pellucida. Note the break in the zona pellucida on the right side of the blastocyst. The zona pellucida–free half of the blastocyst protrudes through the hole in the zona pellucida while the other half (on the left) is still surrounded by it.

Figure 1A.2.10 Zona pellucida removal. Blastocyst-stage embryo before (left) and after (right) zona pellucida removal with acidified Tyrode’s solution. Labels: i, inner cell mass; t, trophectoderm; z, zona pellucida. Change in embryo shape is a sign that the zona pellucida is dissolved.

SUPPORT PROTOCOL 3

REMOVAL OF THE ZONA PELLUCIDA WITH PRONASE Zona pellucida removal with acidified Tyrode’s solution is a rapid process, and it is quite easy for the unskilled experimenter to irreparably damage the embryo. Therefore, some experimenters use the pronase method to remove the zona pellucida, a more time-consuming process that decreases the likelihood of the inadvertent embryo damage. While use of acidified Tyrode’s solution is preferred in the cases when hESC may have potential therapeutic use, because it eliminates the use of animal-derived enzyme (pronase), pronase treatment is in other aspects equivalent to acid hydrolysis with Tyrode’s solution.

Materials KSR embryo culture medium with and without 25 ng/ml bFGF (see recipe) 0.5% (w/v) pronase E (Sigma) in KSR embryo culture medium (see recipe) Embryos in culture (Support Protocol 1) 4-well tissue culture plate with feeder cells (Support Protocol 5) Derivation of hESCs from Intact Blastocysts

6-cm tissue culture dishes with cell culture–treated surface The Stripper micropipettor (MidAtlantic Diagnostics MXL3-STR) and 600-µm tips

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1. Place six separate 50-µl drops of KSR embryo culture medium on a cell culture– treated surface of a sterile 6-cm tissue culture dish (for washing the embryo after pronase treatment). 2. Place two 50-ml drops of 0.5% pronase in the same dish; mark pronase drops to avoid error. 3. Remove embryo from the culture drop under oil using The Stripper micropipettor with a 600-µm tip and transfer into a drop of KSR embryo culture medium. 4. Examine the blastocyst-stage embryo under the microscope and assign a grade (see Fig. 1A.2.8). Do not treat embryos that have initiated hatching with pronase. Instead, transfer them onto feeders in the blastocyst medium and replace the medium with KSR embryo culture medium supplemented with bFGF once the embryo has completely hatched and detached from the zona pellucida.

5. Transfer the embryo into the first drop of pronase for a brief rinse, and then transfer to the second drop of pronase. Transfer dish into incubator and incubate 3 min at 37◦ C. 6. Remove the dish from the incubator and examine the embryo for presence of the zona pellucida. If the zona pellucida is still present, incubate the dish again ∼1 min at 37◦ C. Repeat as many times as necessary. 7. As soon as the zona pellucida is dissolved, quickly transfer the embryo to the first drop of KSR embryo culture medium. 8. Transfer the embryo to the next drop and repeat until the sixth drop. Examine the embryo to ensure that the zona pellucida was completely removed. 9. Place the zona pellucida–free embryo into the well of a 4-well tissue culture plate with feeder cells in 0.5 ml KSR embryo culture medium supplemented with 25 ng/ml bFGF. Place only one embryo into each well. Because each embryo has its own unique genetic material, it is crucial not to mix them.

THAWING EMBRYOS Embryo cryopreservation is a relatively new technique. The first pregnancy from a frozen and thawed human embryo was reported in 1983, and a birth from this source occurred the following year. Of ∼100,000 cases of assisted reproductive technology in the United States in 2000, ∼16% of the cases used frozen and thawed embryos. In 2000, live birth rates per thaw cycle were 18.3% versus 26.6% from the fresh embryo transfer. Theoretically, if there are no temperature variations, the embryos can be frozen indefinitely and still be successfully recovered. Embryos are gradually cooled from the body temperature to −196◦ C in the presence of cryoprotectants (e.g., propanediol) that prevent damage from intracellular ice formation and interact with membranes during their transition from a pliable to a rigid state. Thawing, which means bringing frozen embryos to room temperature, is a quick process, taking less than 2 min. However, the most critical aspect of the process is a slow step-wise exchange of cryoprotectant fluids with culture medium. Once the thawing is completed, the embryo is assessed for cryodamage. If there is no blastomere loss during cryopreservation, cryopreserved embryos are equivalent to fresh embryos. However, some healthy embryos may not survive the stress of freezing and thawing without partial cellular damage and blastomere lysis.

SUPPORT PROTOCOL 4

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Materials Embryos frozen in straws under liquid nitrogen (from IVF center) Embryo Thaw Media Kit containing solutions T1, T2, and T3 (Irvine Scientific) 100 mg/ml human serum albumin solution (HSA; Irvine Scientific) Modified human tubal fluid medium (mHTF; Irvine Scientific) 6-cm tissue culture dish (e.g., Falcon, 3046) The Stripper micropipettor (MidAtlantic Diagnostics MXL3-STR) and appropriate size tips Prepare solutions 1. Verify, using the accompanying documentation, that the straw removed from the liquid nitrogen storage tank contains embryos at the desired stage of development. In vitro fertilization clinics usually freeze embryos at the cleavage stage (day 3), although some may also freeze them at the single-cell, pronuclear stage (day 1) or at the blastocyst stage (day 5 or 6). Thaw media kits are not the same for cleavage- and blastocyst-stage embryos.

2. Bring solutions T1, T2, and T3 from the Embryo Thaw Media Kit to room temperature. 3. Add 12 µl of 100 mg/ml stock solution HSA to 1 ml mHTF. Bring to room temperature. Prepare a second 1-ml aliquot and warm to 37◦ C. Do not use any bottle of HSA which shows evidence of particulate matter, cloudiness, or is not clear pale yellow in color. To avoid problems with contamination, discard any excess medium or HSA stock that remains after the procedure is completed.

Set up thaw plates 4. Put 50 µl of solution T1 into a 6-cm tissue culture dish, and mark the drop as number 1. Embryo thaw solution T1 is a 1.0 M propanediol solution containing 0.2 M sucrose in mHTF supplemented with 12 mg/ml HSA. During the thawing procedure, the cryoprotectant propanediol is removed, and the embryos are rehydrated. Because of its high molecular weight, sucrose does not pass through the plasma membrane, and therefore it is included in the thawing solution to aid in the removal of cryoprotectant via osmosis. Several embryos may be placed into each drop of thawing solution, but to ensure that there is no potential for cross-contamination; only embryos from the same donor should be placed together. The arrangement of the drops of the different solutions on the same or different plates depends on how many embryos are being thawed. More than three drops in one dish might be too close and easily mixed.

5. Put 50 µl solution T2 into the 6-cm tissue culture dish, and mark the drop as number 2. Embryo thaw solution T2 is a 0.5 M propanediol solution containing 0.2 M sucrose in mHTF supplemented with 12 mg/ml HSA.

6. Put 50 µl solution T3 into the 6-cm tissue culture dish, and mark the drop as number 3. Embryo thaw solution T3 is a 0.2 M sucrose in mHTF supplemented with 12 mg/ml HSA.

7. Put 50 µl of HSA/mHTF medium into a separate 6-cm tissue culture dish, and mark the drop as number 4. 8. Fill a 50-ml test tube with sterile water heated to 30◦ C to act as a water bath. Derivation of hESCs from Intact Blastocysts

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Thaw embryo 9. Remove the straw containing frozen embryos from the liquid N2 storage. Hold straw in the air for 30 to 40 sec, then immerse in the 30◦ C water bath for 40 to 60 sec to thaw. While identifying the correct straws, keep them in the liquid nitrogen to prevent temperature increase.

10. Remove the plastic top of the straw. Hold the straw at an angle against a sterile tissue culture dish and push content out, drop by drop. 11. Using the Stripper micropipettor with an appropriate size tip transfer the embryo(s) to drop number 1 with solution T1 and leave 5 min at room temperature. 12. Transfer the embryo(s) to drop number 2 (solution T2) and incubate 5 min at room temperature. 13. Transfer the embryo(s) to drop number 3 (solution T3) and incubate 10 min at room temperature. 14. Transfer the embryo(s) to drop number 4 (mHTF/HSA medium) and incubate 10 min at room temperature. 15. Put 50 µl of prewarmed HSA/mHTF medium into a separate 6-cm tissue culture dish, and mark the drop as number 5. Transfer the embryo(s) to drop number 5 (HSA/HTF medium) prewarmed to 37◦ C and incubate 10 min at 37◦ C. 16. Proceed with embryo culture as described in Support Protocol 1.

PLATING OF FEEDER CELLS Human embryonic stem cells were originally derived on feeder layers of mitotically inactivated mouse embryonic fibroblasts (Thomson et al., 1998). The incorporation of nonhuman sialic N-glycolylneuraminic acid (Neu5Gc) from nonhuman feeder layers and medium by hESC leads to an immune response mediated by natural anti-Neu5Gc antibodies present in most humans (Martin et al., 2005); in cases when there is potential for therapeutic uses of the hESC, it is advantageous to replace mouse embryonic fibroblasts as feeder cells with feeder cells of human origin or, ideally, with a feeder-layer-free culture environment (Ilic, 2006). Among human feeder cells that support not only growth but also derivation of hESC lines, human foreskin (Amit et al., 2003; Hovatta et al., 2003) and placental fibroblasts (Genbacev et al., 2005) are the most easily accessible.

SUPPORT PROTOCOL 5

Materials 0.5% (w/v) gelatin (see recipe) Phosphate-buffered saline (PBS), calcium and magnesium free (Gibco/Invitrogen) Fibroblasts: irradiated and frozen mouse or human cells (see Conner, 2000; Nagy, 2003) Fibroblast feeder medium (see recipe), prewarmed to 37◦ C 15-ml centrifuge tube, sterile 6-well, tissue culture–treated plates (e.g., Corning) or 4-well, tissue culture–treated plated (e.g., Nunc) Additional reagents and equipment for counting cells (Phelan, 2006) Prepare gelatin-coated plates 1. Add 0.5% gelatin to the tissue culture plates (0.5 ml/well of 4-well plate or 2 ml/well of 6-well plate) and incubate at least 2 hr at 37◦ C. Swirl to wet the entire surface of the wells.

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2. Aspirate the gelatin and either use plates immediately or fill wells with PBS and leave in the 37◦ C incubator until use (maximum 3 days).

Thaw and plate irradiated fibroblasts 3. Thaw a cryovial of irradiated fibroblasts at 37◦ C and transfer contents into a sterile, 15-ml centrifuge tube containing 9 ml prewarmed fibroblast feeder medium. 4. Centrifuge the cells 5 min at 700 × g, room temperature. 5. Remove the supernatant and resuspend the cell pellet in fresh fibroblast medium. 6. Count the resuspended cells (see Phelan, 2006) and adjust the cell number according to the plating plan (see step 7 annotation) with additional fibroblast medium. 7. Plate the cells in a volume of fibroblast feeder medium and at cell density adjusted to the surface area of the cell culture plate used (to give 70% to 80% confluency within 3 days). The optimal number of cells should be determined for each lot and type of irradiated cells. When determining the number of cells to be plated, use 1.5 × 104 cells/cm2 as a starting point. For example, plate 2 – 4 × 104 cells in 0.5 ml fibroblast culture medium per well of a 4-well tissue culture plate. Ideally, feeders will be 70% to 80% confluent at the time of embryo plating and not longer than 3 days in culture. However, thawed and plated irradiated fibroblasts may be used as feeders up to 1 week after plating. They are kept in the cell incubator until used. Irradiated fibroblasts are mitotically inactivated, which means that they can only complete a cell division cycle initiated prior to the irradiation, but cannot divide any further. However, in some cases, a few cells might escape mitotic inactivation with mitomycin C or irradiation and proliferate to fill up the well with feeder cells after prolonged culture. Some feeders can support hESC for 4 weeks before they deteriorate, while others are only good for about 2 weeks. How often feeders should be prepared must be determined by the investigators for each type and preparation of feeders used in their laboratories.

8. Change the medium once, 1 day after plating.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Fibroblast feeder medium 360 ml Dulbecco’s modified Eagle medium (DMEM), high glucose (Gibco/ Invitrogen) 90 ml medium 199 (Gibco/Invitrogen) 50 ml heat-inactivated fetal bovine serum (Hyclone): prepared by dividing into 50-ml aliquots and storing up to 1 year at −20◦ C Sterilize by passing through a 0.22-µm l cellulose acetate, low-protein-binding filter (Corning) and store up to 1 month at 4◦ C. Gelatin, 0.5% (w/v) 50 ml 2 % (w/v) gelatin, Type B (Sigma) 150 ml H2 O Sterilize by passing through a 0.22-µm low-protein-binding filter (Corning), divide into 10-ml aliquots, and store up to 1 year at −20◦ C. Thawed 0.5% gelatin can be stored up to 1 week at 4◦ C. Derivation of hESCs from Intact Blastocysts

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Human recombinant basic fibroblast growth factor (bFGF) stock, 10 µg/ml 100 µg human recombinant basic fibroblast growth factor (bFGF; R&D), four 25-µg vials 10 ml diluted human serum albumin solution: prepared by diluting 20 µl 100 mg/ml human serum albumin solution (Irvine Scientific) with 10 ml calciumand magnesium-free PBS (Gibco/Invitrogen) Make up a 10 µg/ml solution of bFGF by dissolving four 25-µg vials of bFGF in a total of 10 ml diluted human Serum Albumin (HSA)/PBS in the original vials. Pool the solutions and sterilize by passing through a 0.2-µm surfactant-free cellulose acetate syringe filter (e.g., Corning 431219) (prefiltered with HSA solution diluted 1/10 in PBS). Divide into 1-ml aliquots and store up to 1 month at −20◦ C or up to 1 year at −80◦ C. Upon thawing, record the thawing date on the tube and store thawed aliquots up to 1 month at 4◦ C. Do not use any bottle of HSA that shows evidence of particulate matter or cloudiness or is not clear pale yellow in color. To avoid problems with contamination, discard any excess medium or HSA stock that remains after the procedure is completed.

KSR embryo culture medium, with and without 25 ng/ml bFGF 400 ml Knockout Dulbecco’s modified Eagle medium (e.g., Gibco/Invitrogen) 100 ml Knockout Serum Replacement (Gibco/Invitrogen) 5 ml 200 mM L-glutamine (Gibco/Invitrogen) 5 ml 10 mM modified Eagle medium nonessential amino acids solution, 100× stock (Gibco/Invitrogen) 1 ml 0.1 mM 2-mercaptoethanol Sterilize by passing through a 0.22-µm cellulose acetate, low-protein-binding filter unit (Corning). Store up to 1 month at 4◦ C. When required, add human recombinant bFGF (see recipe) to an aliquot of KSR embryo culture medium in a sterile tube to a final concentration of 25 ng/ml. Store up to 24 hours at 4◦ C.

2-Mercapoethanol stock, 0.1 mM Combine 53 µl 99% 2-mercapoethanol (Sigma) with water to a final volume of 15 ml. Sterilize by passing through a 0.2-µm regenerated cellulose syringe filter (Corning), and divide into 1.5-ml aliquots. Store up to 6 months at –20◦ C.

COMMENTARY Background Information Embryonic stem cells (ESC) originate from the pre-implantation mammalian embryo. As it travels down the oviduct, a fertilized oocyte (or zygote) divides to generate a 16- and 32cell morula (Johnson and McConnell, 2004). With subsequent cell divisions, a blastocoel cavity forms in the center of the morula and embryonic cells differentiate into two morphologically distinct populations within the blastocyst: an outer layer of cells comprising the trophectoderm, which will form placenta, and the inner cell mass (ICM) that will give rise to the fetus. The cells from the ICM give rise to ESC in culture. However, the pluripotent cell population that exists for a short time within ICM

of the developing blastocyst is most likely not identical to the derived ESC. During derivation, ESC undergo epigenetic changes to adjust to cell culture conditions and therefore acquire certain characteristics which separate them from the embryonic cells from which they originate (see Krtolica and Genbacev, 2007). However, hESCs share with embryonic ICM cells a pluripotent capacity and capability of self-renewal (Amit et al., 2000; Draper and Fox, 2003). During ESC differentiation in culture, as well as embryonic differentiation in vivo, heterochromatin formation selectively suppresses gene expression, resulting in a loss of pluripotent capacity (Rasmussen, 2003). It is interesting to note that while the differentiation

Isolation of Embryonic Stem Cells

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potential of the ICM cells in vivo is not equivalent to a totipotent zygote—they do not form placenta and some other extraembryonic tissues—ESC in culture may have somewhat extended differentiation capacity and can give rise to trophectoderm-like cells (Xu et al., 2002). Unlike the majority of somatic cells which undergo telomere shortening with each cell division and as a result have finite life span that ends with senescent arrest (Krtolica and Campisi, 2002), ESC express telomerase, a reverse transcriptase that adds telomeric DNA to chromosome ends thus preventing telomere shortening and growth arrest (Verfaillie et al., 2002; Carpenter et al., 2003). In this way, ESC maintain their telomere length at 8 to 12 kb and are capable of unlimited selfrenewal (Verfaillie et al., 2002). When grown in culture, they exhibit a virtually indefinite replicative lifespan—some ESC lines have been propagated for years without any signs of slowing down. Although reported derivation rates vary significantly between the investigators, there does not appear to be consistent difference in the efficiency of derivation between those who use the isolated ICM and those who start with the intact blastocysts. However, using intact blastocysts provides some advantages: It eliminates technically challenging step of ICM isolation which requires either micromanipulator for the mechanical/laser dissection or immunosurgery. It abrogates the exposure of the embryos to animal-derived complement that is used to destroy trophectoderm cells during immunosurgery, a most common procedure for the isolation of the ICM. This may be advantageous in case derived ESC are intended for clinical use. It avoids risk of damaging the ICM during removal of trophectoderm. It enables use of underdeveloped blastocysts in which ICM may not be clearly visible. That said, some groups reported high efficiency of ESC derivation using isolated ICM, and there is no question that both methods can yield ESC of similar characteristics.

Critical Parameters

Derivation of hESCs from Intact Blastocysts

All tissue culture must be performed in Class II biological safety cabinets or laminar airflow workstations. All reagents and media must be sterilized (except for presterilized em-

bryo media) by passing through 0.22-µm filters and should be discarded after their expiration date. Embryo transfer and removal of the zona pellucida should be performed in the shortest possible time to reduce stress and exposure to nonoptimal culture conditions. Even if all procedures are performed correctly, the embryo may not give rise to hESC. The success of hESC derivation ultimately depends on two parameters: quality of the embryos and quality of the feeder cells. In the authors’ experience, embryos with larger and well defined inner cell masses are more likely to give a rise to an hESC line. It is essential that feeders are freshly plated (1 to 3 days before use) and at the right density. It is also recommended that feeder cells used for derivation be from the passages/population doublings within the first 30% to 50% of their lifespan (i.e., between passages 7 and 12 if split 1:2 for human placental fibroblasts, passages 4 to 5 for mouse embryo fibroblasts, and <18 passages for foreskin fibroblasts). However, earlier passages may yield more efficient derivation); earlier passages are better.

Troubleshooting To reduce technical problems, staff involved in hESC derivation should be experienced in handling cleavage- and blastocyststage embryos. If not, it is recommended that they practice first on mouse, bovine, or hamster eggs/embryos. If removal of the zona pellucida is a challenge, one can wait for spontaneous hatching. In this case, the expanded blastocyst-stage embryo should be transferred onto feeders in the blastocyst medium. The medium should be replaced with KSR medium supplemented with bFGF once the embryo has completely hatched and detached from the zona pellucida. Because spontaneous hatching may or may not occur, and it happens only in the fully expanded highgrade blastocyst-stage embryos, relying only on hatched blastocysts would markedly reduce number of embryos available for hESC derivation. In case grade 1 and 2 embryos with A or B inner cell mass consistently fail to give rise to an ESC line, it is recommended that the feeders used should be replaced by earlier passage and/or different feeder cell line. If the growth of feeder cells is prominent in ESC cultures, a higher dose of irradiation or mitomycin C should be used for their mitotic inactivation.

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Anticipated Results Good quality cleavage stage (day 3) embryos (grades 1 and 2) are likely to develop into blastocysts with an efficiency of 40% to 50%. For lower grade embryos the efficiency of reaching the blastocyst stage is significantly lower. It is anticipated that between 20% and 50% of blastocysts will be able to give rise to embryonic stem cell colonies. The efficiency of derivation depends in large part on the quality of the blastocyst and the number and quality of inner cell mass cells in particular. Typically, blastocysts with the inner cell mass grade A or B are more efficient in yielding the ESC line. Some of the derived ESC lines may develop abnormal karyotypes, and it is therefore recommended that the initial freezing of ESC be performed as early as possible. The karyotype of each newly derived ESC line should be analyzed by G-banding after initial freezing and every 5 to 10 passages thereafter.

mercaptoethanol stock and dividing in aliquots takes about 30 min for each. Placing embryos after zona pellucida removal onto feeders should not take more than few minutes. Attachment of embryos onto feeders takes between 24 and 48 hr. The derivation process from the moment embryos are placed onto feeders until the first visible embryonic stem cell colony outgrowth arises takes usually between 2 and 4 weeks. It takes additional 4 to 12 weeks to expand cells enough to be able to freeze them. Plating feeder cells To prepare fibroblast growing medium and gelatin stock takes about 30 min for each. Gelatin coating of tissue culture plates and thawing and plating of irradiated fibroblasts takes about 3 hr or more, depending on the number of plates being coated.

Literature Cited Time Considerations Thawing embryos The procedure takes approximately 2 hr, including media preparation. Culturing embryos Time needed depends on the number of drops and number of embryos. Preparation of embryo culture medium in droplets and covering with oil requires 1 to 2 min per one 6-cm tissue culture dish. Drops need to be pre-incubated for at least 1 hr prior to their use. Transfer of embryos from one drop to another is fairly quick, and it should take less than a minute per drop. After thawing, it takes 2 days for pronuclear-stage (day 1) embryos to reach cleavage stage. It takes 2 to 3 days for cleavage-stage (day 3) embryos to develop into blastocysts. Removal of the zona pellucida The process of zona pellucida removal will depend on the number of embryos, competence of the operator, and technique used. Approximately 10 min should be sufficient to remove the zona pellucida from one embryo using acidified Tyrode’s solution, including assigning the grade. Using pronase instead of acidified Tyrode’s solution will prolong zona pellucida removal by 3 to 5 min. hESC derivation Preparation of the KSR embryo culture medium takes about 30 min including thawing of premade frozen aliquots of various constituents. Preparation of bFGF and 2-

Amit, M., Carpenter, M.K., Inokuma, M.S., Chiu, C.P., Harris, C.P., Waknitz, M.A., ItskovitzEldor, J., and Thomson, J.A. 2000. Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227:271278. Amit, M., Margulets, V., Segev, H., Shariki, K., Laevsky, I., Coleman, R., and Itskovitz-Eldor, J. 2003. Human feeder layers for human embryonic stem cells. Biol. Reprod. 68:2150-2156. Carpenter, M.K., Rosler, E., and Rao, M.S. 2003. Characterization and differentiation of human embryonic stem cells. Cloning Stem Cells 5:7988. Conner, D.A. 2000. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Mol. Biol. 51:23.2.1-23.2.7. Draper, J.S. and Fox, V. 2003. Human embryonic stem cells: Multilineage differentiation and mechanisms of self-renewal. Arch. Med. Res. 34:558-564. Genbacev, O., Krtolica, A., Zdravkovic, T., Brunette, E., Powell, S., Nath, A., Caceres, E., McMaster, M., McDonagh, S., Li, Y., Mandalam, R., Lebkowski, J., and Fisher, S.J. 2005. Serum-free derivation of human embryonic stem cell lines on human placental fibroblast feeders. Fertil. Steril. 83:1517-1529. Hovatta, O., Mikkola, M., Gertow, K., Stromberg, A.M., Inzunza, J., Hreinsson, J., Rozell, B., Blennow, E., Andang, M., and Arhlund-Richter, L. 2003. A culture system using human foreskin fibroblasts as feeder cells allows production of human embryonic stem cells. Hum. Reprod. 18:1404-1409. Ilic, D. 2006. Culture of human embryonic stem cells and extracellular matrix microenvironment. Regener. Med. 1:95-101.

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Johnson, M.H. and McConnell, J.M. 2004. Lineage allocation and cell polarity during mouse embryogenesis. Semin. Cell Dev. Biol. 15:583-597.

man embryos in alpha modification of Eagle’s medium under low oxygen tension and low illumination. Fertil. Steril. 62:1022-1027.

Krtolica, A. and Campisi, J. 2002. Cancer and aging: A model for the cancer promoting effects of the aging stroma. Int. J. Biochem. Cell Biol. 34:1401-1414.

Phelan, M.C., 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.8.

Krtolica, A., and Genbacev, O. 2007 Cell polarity, pluripotency and differentiation. In Stem Cells in Human Reproduction (C. Simon and A. Pellicer, eds.) pp. 183-188. Informa Healthcare, London. Martin, M.J., Muotri, A., Gage, F., and Varki, A. 2005. Human embryonic stem cells express immunogenic nonhuman sialic acid. Nat. Med. 11:228-232 Nagy, A., Gertsenstein, M., and Vintersten, K. 2003. Manipulating the Mouse Embryo: A Laboratory Manual, 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Noda, Y., Goto, Y., Umaoka, Y., Shiotani, M., Nakayama, T., and Mori, T. 1994. Culture of hu-

Rasmussen, T.P. 2003. Embryonic stem cell differentiation: A chromatin perspective. Reprod. Biol. Endocrinol. 1:100. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Verfaillie, C.M., Pera, M.F., and Lansdorp, P.M. 2002. Stem cells: Hype and reality. Hematology Am. Soc. Hematol. Educ. Program 369-391. Xu, R.H., Chen, X., Li, D.S., Li, R., Addicks, G.C., Glennon, C., Zwaka, T.P., and Thomson, J.A. 2002. BMP4 initiates human embryonic stem cell differentiation to trophoblast. Nat. Biotechnol. 20:1261-1264.

Derivation of hESCs from Intact Blastocysts

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Reprogramming Primordial Germ Cells (PGC) to Embryonic Germ (EG) Cells

UNIT 1A.3

Gabriela Durcova-Hills1 and Azim Surani1 1

The Wellcome Trust/Cancer Research UK Gurdon Institute of Cancer and Developmental Biology, Cambridge, United Kingdom

ABSTRACT In this unit we describe the derivation of pluripotent embryonic germ (EG) cells from mouse primordial germ cells (PGCs) isolated from both 8.5- and 11.5-days post-coitum (dpc) embryos. Once EG cells are derived we explain how to propagate and characterize the cell lines. We introduce readers to PGCs and explain differences between PGCs and their in vitro derivatives EG cells. Finally, we also compare mouse EG cells with ES cells. This unit will be of great interest to anyone interested in PGCs or studying the behavior of cultured PGCs or the derivation of new EG cell lines. Curr. Protoc. Stem Cell C 2008 by John Wiley & Sons, Inc. Biol. 5:1A.3.1-1A.3.20.
Keywords: primordial germ cells r embryonic germ cells r mouse r reprogramming r pluripotency

INTRODUCTION Germ cells are the source of totipotency but they are also some of the most differentiated cell types. Primordial germ cells (PGCs), embryonic precursors of germ cells, are not pluripotent; that is, they do not self-renew and do not make chimeras. However, they can give rise to pluripotent stem cells, which make chimeras readily. This reversal of the PGCs’ differentiated state to a state of developmental pluripotency is defined here as “reprogramming.” In vivo, transcriptional repression and chromatin remodeling block PGCs from acquiring an overtly pluripotent stem cell phenotype during their differentiation into mature gametes despite their maintenance of Oct-3/4, Sox-2, and nanog expression. This also suggests that PGC development involves a careful balance between inhibition of somatic differentiation genes and maintenance of a “pluripotency program” present in embryonic cells. However, cultured PGCs exposed to appropriate factors are released from the repression and form pluripotent stem cells termed embryonic germ (EG) cells. EG cells, in contrast to PGCs, when exposed to appropriate culture conditions have the capacity to self-renew and differentiate into a variety of cell types. EG cells resemble embryonic stem (ES) cells closely in the morphology of colonies, the culture conditions required for their self-renewal, and the expression of both cell surface and intracellular markers. EG cells, however, differ from ES cells, pluripotent stem cells derived from pre-implantation embryo, with respect to the methylation patterns of their DNA reflecting the erasure of genomic imprints from certain imprinted genes. EG cells derived from 8.5 dpc PGCs make chimeras including germ line. However, many of the chimeras made with EG cells derived from 11.5 dpc PGCs exhibited skeletal malformations. This unit describes generation of EG cells from cultured mouse PGCs isolated from different gestational stages, followed by description of propagation of EG cell lines and characterization of newly established cell lines. The protocols in this unit are designed to provide a basis for the derivation of mouse EG cell lines from 8.5 or 11.5 dpc PGCs (see Basic Protocol) followed by PGC purification from surrounding somatic cells (see Support Protocol 1). Description of Sl4 -m220 feeder

Current Protocols in Stem Cell Biology 1A.3.1-1A.3.20 Published online April 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01a03s5 C 2008 John Wiley & Sons, Inc. Copyright


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cell culture and their mitotic inactivation with mitomycin C (Support Protocol 2) are presented. Characterization of newly derived EG cell lines including PCR sexing (Support Protocol 3), AP staining (Support Protocol 4), and expression of Oct-3/4 and SSEA-1 (Support Protocol 5) are also described. NOTE: All procedures should be performed under sterile conditions. All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All tissue cultures are prepared in a Class II Biological Hazard laminar-flow hood. BASIC PROTOCOL

DERIVING, CULTURING, AND FREEZING MOUSE EMBRYONIC GERM CELLS Mouse embryonic germ (EG) cells are derived from PGCs cultured under appropriate conditions. Here we describe the basic culture technique for derivation of EG cell lines from PGCs isolated from 8.5- and 11.5-dpc embryos. During the first 10 days of culture some PGCs are reprogrammed into EG cells when they form large multicellular colonies resembling EG cell colonies and, moreover, cells coming from these colonies make chimeras (Durcova-Hills et al., 2006). NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Committee for Ethical Animal Care and Use (IACUC) and must conform to government regulations for the care and use of laboratory animals.

Materials 8.5 or 11.5 dpc pregnant MF1 female crossed with 129 male Dissecting medium [make 1% FBS in CMF-PBS (Invitrogen, pH 7.2, 1×, CaCl2 and MgCl2 -free, cat. no. 20012)], sterile 0.05% Trypsin/EDTA (Invitrogen, cat. no. 25300) PGC growth medium (see recipe), sterile 4-well plates with Sl4 -m220 cells, mitotically inactivated (see Support Protocol 2) Mouse embryonic fibroblast (MEF) medium (see recipe) Ca2+ /Mg2+ -free phosphate-buffered saline (CMF-PBS; Invitrogen), sterile EG cell growth medium (see recipe), sterile 35-mm tissue culture dishes seeded with mitotically inactive MEFs (UNIT 1C.3) DMEM Freezing solution (see recipe), sterile Liquid nitrogen 10-cm culture dishes for dissection of fetuses and PGCs-containing tissues Forceps with sharp tips (sterile) for isolation of tissues containing PGCs Dissecting stereomicroscope and inverted microscope 1.0-ml and 100-µl micropipettor tips, sterile 1.5-ml microcentrifuge tubes for collecting tissues, sterile 5-cm tissue culture dish 37◦ C water bath 4-well culture plates to culture PGCs or expand EG cells Pulled-glass mouth pipet 15-ml tubes, sterile Cryotubes −80◦ C freezer

Reprogramming PGC to EG Cells

Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006)

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Figure 1A.3.1 Localization of mouse PGCs. At 8.5 dpc embryo, PGCs are localized at the base of allantois (A). At 11.5 dpc embryo, PGCs are localized in genital ridges, future gonads (B). Mesonephros is attached to the genital ridge and must be removed before preparing PGC suspension (C).

Collect PGC-containing tissues Collect tissues containing 8.5 dpc PGCs 1a. Sacrifice an 8.5-day pregnant female mouse by cervical dislocation (Donovan and Brown, 2006). Using forceps, dissect embryos away from decidua and extraembryonic membranes, in dissecting medium. Other approved methods of euthanasia are carbon dioxide inhalation, barbiturate overdose, or anesthetic overdose. Euthanasia must be carried out by properly trained personnel. Consult your animal care committee for a method used in your institution.

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2a. Using sharp forceps, remove the posterior part of the embryo, starting from the end of the primitive streak and continuing to the base of the allantois. Approximately 100 to 150 PGCs are located at the base of the allantois and we call this region a PGC-containing fragment (Fig. 1A.3.1A).

3a. Collect several PGC-containing fragments with a 1-ml micropipettor tip into a 1.5-ml microcentrifuge tube containing 0.5 ml dissecting medium. The whole dissection procedure should not exceed 1 hr. If longer times are required the microcentrifuge tube with samples should be kept on ice. PGC-containing fragments are pooled from at least eight embryos into a microcentrifuge tube. However, if derivation of EG cell lines from individual embryos is required, the PGC-containing fragments are transferred individually into 1.5-ml microcentrifuge tubes and processed for making cell suspensions.

Collect tissues containing PGCs from 11.5 dpc embryos 1b. Sacrifice an 11.5-day pregnant female mouse (see above) and dissect the embryos away from their extraembryonic membranes. Cut the head off from each embryo in the dissecting medium using sharp forceps. Make a cut along the ventral midline of embryos. Remove all internal organs. 2b. Remove the urogenital ridges (genital ridges with mesonephros) which lie on the dorsal wall of the embryo fragment by the dorsal aorta. Hold the dorsal aorta at the anterior part of the embryo and peel off both the aorta and the urogenital ridges. Collect the urogenital ridges in a 5-cm tissue culture dish filled with dissecting medium (Fig. 1A.3.1B). 3b. Separate the genital ridges from the mesonephros using a needle or sharp forceps (Fig. 1A.3.1C). Subsequently, collect the genital ridges in a 1.5-ml microcentrifuge tube containing 0.5 ml dissecting medium. Genital ridges are pooled from at least five embryos or genital ridges from individual embryos can be collected and processed separately. If it is desirable for further experiments, PGCs can be purified from surrounding somatic cells using magnetic beads (see Support Protocol 1). It is important to separate mesonephros from genital ridges. If not, somatic cells will outgrow the cultured PGCs soon after placing them into culture. Because different embryos from the same mother can vary in their stage of development, it is advisable to classify them and collect them according to the morphology of their hind limb bud. For 11.5 dpc genital ridges the limb bud is symmetric and has a circular outline or an outline of digits is just visible.

Culture PGCs leading to the derivation of EG cell line 4. Centrifuge collected tissue samples 5 min at 250 × g, room temperature. 5. Add 100 to 200 µl of trypsin/EDTA (the volume depends on the amount of tissue) and incubate 5 to 10 min in a 37◦ C water bath. 6. Obtain cell suspension by gently pipetting tissues with a 100-µl micropipettor tip. Add 800 µl of PGC growth medium to trypsinized cell suspension and mix well to inactivate trypsin. Cell suspension is obtained by pipetting with either a 100-µl micropipettor tip when PGC-containing tissues are isolated from 8.5 dpc embryos or a 1-ml micropipettor tip when genital ridges are isolated from 11.5 dpc embryos.

Reprogramming PGC to EG Cells

7. Centrifuge 5 min at 250 × g, room temperature. Resuspend the pellet in 500 to 1000 µl of the PGC growth medium.

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Figure 1A.3.2 PGCs cultured in LIF, FGF-2, and SCF after 8 to 10 days give rise to a multicellular colony containing 200 to 300 cells (A). The colony is further expanded generating many colonies (B) expressing AP (C).

8. Plate cell suspension onto mitotically inactivated Sl4 -m220-containing wells of 4-well culture dishes (see Support Protocol 1) containing 0.5 ml of PGC growth medium supplemented with LIF and FGF-2. Plate approximately half of the 8.5-dpc PGC-containing fragment or 0.2 to 0.25 of one 11.5-dpc genital ridge into each well of a 4-well dish.

9. Culture in an incubator at 37◦ C, 5% CO2 in air. After 2 days, change the PGC growth medium with medium freshly supplemented with both LIF (1200 U in 1 ml medium) and FGF-2 (25 ng in 1 ml medium). Then change the medium every day or every other day as needed. When the medium is orange/yellow in color it indicates that the pH of the medium is too acidic (low pH) as a result of cell metabolism and the medium should be replaced with fresh medium supplemented with LIF and FGF-2.

10. Culture 8 to 10 days until small colonies are observed. 11. Prepare mitotically inactive primary mouse embryonic fibroblasts (MEFs) on 4-well culture plates in 0.5 ml (MEF medium) at a density of 2 × 105 cells/ml. 12. Culture 2 further days to allow colonies to enlarge in size (200 to 300 cells per colony; Fig. 1A.3.2A) so they are ready to be transferred onto the MEFs. 13. Pick colonies from culture plates with a pulled-glass mouth pipet, immediately transfer to a well containing 0.5 ml CMF-PBS in a 4-well plate. Wash every colony twice, each time in 0.5 ml CMF-PBS. 14. Transfer to a well containing 0.5 ml trypsin/EDTA in a 4-well plate. Incubate for 5 to 10 min at 37◦ C, until individual cells within a colony can be identified. There are usually 2 to 3 colonies per one well in a 4-well plate. PGCs reprogrammed into pluripotent stem cells form large multicellular colonies that by morphology resemble EG colonies, which usually are composed of 200 to 300 cells. Nonreprogrammed PGCs form only small colonies that do not resemble an EG colony.

15. Transfer one of the colonies from one well using a glass pipet into a well of a 4-well plate containing both the mitotically inactive MEF and 0.5 ml of the EG cell growth medium supplemented only with LIF (1000 U in 1 ml culture medium). Embryonic and Extraembryonic Stem Cells

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16. Gently disaggregate each colony into small clumps by pulling and pushing the colony up and down in the glass mouth pipet (trituration). Do not try to disaggregate colony into single cells because many cells could be damaged. Use mitotically inactivated MEFs for further propagation of newly derived EG cells.

17. Culture in an incubator at 37◦ C, 5% CO2 in air, for a few days. After 1 to 2 days the small colonies will start growing on MEF feeders.

18. After 2 additional days, examine colonies, and select colonies which will be used for further culture. Morphology of colonies resembling EG cell colony are used as selection criteria.

Passage colonies 19. Transfer selected colonies (10 to 20) by glass mouth pipet into 0.5 ml CMF-PBS, followed by washing with 0.5 ml CMF-PBS and incubating in 0.5 ml trypsin/EDTA to obtain a cell suspension of 2- to 5-cell clumps. 20. Transfer cell suspension onto mitotically inactivated MEF feeders containing EG cell growth medium in one well of a 4-well plate. Make sure that every colony was disaggregated into 2- to 5-cell clumps. If very large clumps (containing 20 or more cells) are observed remove them from the culture using the glass pipet.

21. After an additional 3 to 4 days of culture observe colonies, which will be 50 to 100 cells. 22. Aspirate the culture medium, wash each well with 0.5 ml CMF-PBS and add 0.5 ml 0.05% trypsin/EDTA. Incubate for 5 to 10 min at 37◦ C and make a single-cell suspension by pipetting with a 100-µl micropipettor tip. 23. Inactivate the trypsin by adding 0.5 ml of EG cell growth medium. 24. Transfer the cell suspension into a 1.5-ml microcentrifuge tube and centrifuge 10 min at 250 × g, room temperature, to obtain a pellet. 25. Resuspend the pellet in 2 ml EG cell culture medium. Transfer the cell suspension into 35-mm dish, on mitotically inactive MEF feeders Resume incubation. If only a few colonies are observed, transfer them into a new well in a 4-well dish. After 3 to 4 days many colonies will be growing in the culture dish (Fig. 1A.3.2B).

26. Sub-culture 10 to 20 colonies into a 35-mm culture dish as described before (Fig. 1A.3.2A). Transfer of EG cells into a 35-mm culture dish is counted as the first passage.

Freeze newly derived EG cells 27. Passage cells 1 or 2 more times to get enough EG cells to freeze down in a few cryotubes in a liquid nitrogen storage tank. Record it in cell line recording book. For EG cell line derivation it is preferable to isolate and propagate an individual multicellular colony, which eventually will give rise to an EG cell line, rather than pool several colonies into one sample.

Manipulations with established cell line

Reprogramming PGC to EG Cells

Grow EG cells 28. Grow EG cells in 35-mm culture dishes or 6-well plate on mitotically inactive MEF cells in the EG cell growth medium supplemented only with LIF. Inspect the culture

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every day and change the medium daily or every other day. After 2 to 3 days or when near confluency, cells are ready to be subcultured or frozen. We usually grow one EG cell line in 3 to 4 wells of a 6-well plate. Do not allow the culture medium during the EG cells growth to turn yellowish (indicating a low pH) because it might induce differentiation of EG cells. EG cell colonies during this period enlarge in size while keeping an undifferentiated morphology. The undifferentiated EG cell colony usually has a round or oval shape with a distinct border around the colony in which individual cells are tightly compacted.

Passage EG cells 29. Grow EG cells on MEF feeders for 2 to 3 days in 35-mm culture dishes or wells of a 6-well plate in EG cell medium supplemented with LIF. 30. Wash wells with 1 ml CMF-PBS, add 0.5 ml trypsin and incubate in the incubator for 5 to 10 min. 31. Prepare a single-cell suspension by pipetting the trypsinized cells with a 1-ml micropipettor tip. 32. Add 1 ml of EG cell growth medium to neutralize trypsin. Transfer the cell suspension into a 15-ml tube containing 8 ml of DMEM. 33. Centrifuge 5 min at 800 × g, room temperature. 34. Resuspend the pellet in 6 to 8 ml of EG cell growth medium (1:4 dilution). 35. Plate 2 ml of EG cell suspension per one 35-mm culture dish or a well in a 6-well plate. Culture cells at 37◦ C in an incubator.

Freeze EG cells 36. EG cells that are subconfluent and in growth phase are ready to be frozen down. Remove the EG cell growth medium and wash cultures with 1 ml CMF-PBS. EG cells are subconfluent when they cover ∼85% to 90% of culture dish.

37. Make a single-cell suspension by trypsin/EDTA treatment, then pellet the cells as described above (steps 30 to 33). 38. Resuspend the pellet in cold freezing solution. Use 1 ml of freezing solution for EG cells collected from one 35-mm culture dish or one well in 6-well plate. 39. Transfer cells in freezing solution to cryotubes and label with cell name, passage number, and date. If the EG cell line was grown in four 35-mm culture dishes or wells, resuspend the pellet obtained by trypsinization in 4 ml of freezing solution, then aliquot into four cryotubes.

40. Place cryotubes overnight in an −80◦ C freezer. 41. Next day transfer cryotubes to liquid nitrogen and record position in cell database file.

Thaw EG cells 42. Thaw frozen EG cells quickly by placing the cryotube into a water bath at 37◦ C. Special safety precautions should be taken when retrieving vials from liquid nitrogen (LN2 ) storage. Protect eyes by wearing protective glasses. The potential hazard exists that LN2 has entered the vial and rapid expansion of the LN2 could cause an explosion and dissemination of the contents of the vial.

43. Transfer cell suspension into a 15-ml tube containing 9 ml DMEM.

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44. Centrifuge 5 min at 800 × g, room temperature. 45. Resuspend the cell pellet in 8 ml of the EG cell growth medium (1:4 dilution) and seed 2 ml of EG cell suspension onto mitotically inactive MEF cells in a 35-mm culture dish, so that EG cells coming from one cryotube will be divided into four 35-mm culture dishes. 46. Place culture into the CO2 incubator. Next day examine the cultures for small colonies of growing EG cells. Change the growth medium to remove dead, floating cells. 47. Feed cultures every day and check under a microscope for colony appearance and potential contaminations. It takes ∼3 to 4 days to obtain cells at subconfluent density and in a growing phase. At this stage of growth they should be used either for immunofluorescence staining, or for making chimeras. For Southern or immunoblotting, cells must be further expanded into 10-cm culture dishes and then used for isolation of genomic DNA or proteins. SUPPORT PROTOCOL 1

PURIFICATION OF 11.5 dpc PGCs FROM SURROUNDING SOMATIC CELLS BY APPLYING MAGNETIC BEADS For some applications it may be desirable to purify PGCs from somatic cells. Here we present a fast, reliable protocol for PGC purification using magnetic beads and a magnetic field. Using this technique the achieved PGC purity is ∼90% to 95% and viability of sorted cells is 95%. If PGCs of the same sex are desirable, first determine the sex of each embryo by PCR (see Support Protocol 3) and then collect genital ridges from embryos of the same sex. This method is adapted from that published by Pesce and De Felici (1995).

Additional Materials (see Basic Protocol) Cell sorting medium (see recipe), cold SSEA-1 antibody (Developmental Studies Hybridoma Bank, The University of Iowa, or Abcam) AP staining solution (see Support Protocol 4) MiniMACS Starting Kit (Miltenyi Biotec) containing: 1 MiniMACS Separating Unit 1 MACS MultiStand 25 MS Columns 1 ml unit of MACS MicroBeads (rat-anti mouse IgM) Plate shaker in a cold room Additional reagents and solutions for counting cells using a hemacytometer (Phelan, 2006) Digest genital ridges 1. Collect ten 11.5 dpc genital ridges in 1.5-ml microcentrifuge tube. Wash them twice with 1 ml of CMF-PBS to remove any traces of FBS. FBS inhibits the activity of trypsin. ◦

2. Incubate the genital ridges in 0.250 ml 0.05% trypsin/EDTA for 5 to 10 min at 37 C in a water bath. 3. Pipet up and down five times with a 1-ml micropipettor tip attached to a pipettor and then a 100-µl micropipettor tip to obtain a single-cell suspension.

Reprogramming PGC to EG Cells

It is important to obtain a single-cell suspension in order to achieve a high purity. Therefore transfer the cell suspension into a well in a 4-well dish and check the suspension under the microscope. If you see any clumps of cells remove them by using a glass mouthpipet or a 100-µl micropipettor tip. Then transfer the cell suspension back into a 1.5-ml microcentrifuge tube.

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4. Add 500 µl cold cell sorting medium. Centrifuge 5 min at 250 × g, 6◦ C. 5. Resuspend the pellet in 500 µl of the cold sorting medium to obtain a single-cell suspension.

Expose cell to antibody 6. Add 50 µl of SSEA-1 antibody and incubate on a plate shaker for 45 min at 4◦ C in order to allow SSEA-1 antibodies to bind to the antigen present on PGCs cell surface. The amount of primary antibody necessary for successful separation must be tested with each new lot of antibody and appropriately diluted if needed.

7. Gently centrifuge 3 min at 200 × g, 6◦ C. Gently resuspend the pellet in 500 µl of cold sorting medium using a 1-ml micropipettor tip attached to a pipettor.

Bind beads 8. Add 20 µl of secondary antibody–coated microbeads (microbeads conjugated with rat-anti mouse IgM) and incubate on a plate shaker for 15 to 25 min at 4◦ C as before. Check if the cell suspension is gently moving so that cells will not form small aggregates during the incubation time.

9. Add 0.5 ml of cold sorting medium and gently pipet with a 1-ml micropipettor tip attached to a pipettor.

Separate PGCs from somatic cells 10. Prewash the column with 1 ml of the cold sorting medium and subsequently pipet the magnetic-bead coated cell suspension (1 ml) on top of the prewashed column. Do not allow the column to dry out. Work in a laminar flow hood if purified PGCs will be cultured.

11. Collect the negative fraction, i.e., cells which were not magnetic-bead stained. This fraction contains somatic cells of genital ridges. If the somatic cells are not needed for further work they are discarded.

12. Wash the column with 0.5 to 1 ml of the cold sorting medium. Then remove the column from the magnet and flush magnetically stained cells into a 1.5-ml microcentrifuge tube with 1 ml of the cold sorting medium by using the plunger. Collect as a positive fraction, i.e., PGCs which were magnetic-bead stained.

Assess the purity of the preparation 13. To determine the purity of the positive fraction, stain 50 µl from the positive fraction with alkaline phosphatase (AP) solution (see Support Protocol 3). 14. Count the number of AP-positive and AP-negative cells with a hemacytometer (Phelan, 2006) and calculate the purity. 15. Seed the remaining cells from the positive fraction onto dishes/plates with Sl4 -m220 feeders in medium used for derivation of EG cell lines.

PREPARATION OF MITOTICALLY INACTIVATED Sl4 -m220 FEEDER CELLS

SUPPORT PROTOCOL 2

The Sl4 -m220 cell line is used for PGCs reprogramming into EG cells which occurs over the first 10 days of culture. Once PGCs, co-cultured with Sl4 -m220 cells in the presence of LIF and FGF-2 in medium, form a large colony resembling an EG cell colony, they are further propagated on mouse embryonic fibroblast (MEF) cells. Sl4 m220 cells express the membrane-bound form of SCF. Sl4 -m220 cells can be obtained from Dr. Matsui from the Cell Resource Center for Biomedical Research, Institute of

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Development, Aging and Cancer, Tohoku University, Japan. Sl4 -m220 can be replaced by STO cell line, which is commercially available from European Collection of Cell Culture (http://www.ecacc.org.uk).

Materials Sl4 -m220 cells, frozen Sl4 -m220 growth medium (see recipe), sterile Mitomycin C solution (see recipe), sterile Ca2+ /Mg2+ -free phosphate-buffered saline (CMF-PBS; Invitrogen), sterile 0.05% Trypsin/EDTA PGC growth medium (see recipe) 10-cm petri dishes, gelatinized (see recipe) 1-ml micropipettor tip attached to a pipettor 15-ml tubes 4-well culture dishes, gelatinized (see recipe) Additional reagents and equipment for counting cells (UNIT 1C.3) Plate Sl4-m220 cells 1. Plate one cryotube of Sl4 -m220 cells onto five pregelatinized 10-cm culture dishes in 10 ml Sl4 -m220 growth medium 4 days before starting the PGC culture. 2. Replace medium with 10 ml fresh growth medium after 2 days of culture. After an additional 1 day of culture, Sl4 -m220 cells should reach subconfluency.

3. On day 4, replace the growth medium of subconfluent cultures in all dishes with 5 ml mitomycin C–containing medium. One 10-cm dish will contain 5 ml of Sl4 -m220 growth medium and 12.5 µl of mitomycin C from the stock solution. Sl4 -m220 cells are subconfluent when they occupy 95% of 10-cm dish space.

4. Incubate for 2 hr. 5. Remove the mitomycin C–containing medium and rinse the cells three times, each time with 5 ml CMF-PBS to remove any traces of mitomycin C.

Trypsinize cells 6. Incubate cells with 1 ml 0.05% trypsin/EDTA for 5 min at room temperature. Pipet cells with a 1-ml micropipettor tip attached to a pipettor to harvest a single-cell suspension. 7. Collect trypsinized cells in a 15-ml conical tube containing 5 ml of the Sl4 -m220 growth medium to inactivate and dilute the trypsin solution. 8. Centrifuge 5 min at 800 × g, room temperature, and remove solution. 9. Resuspend Sl4 -m220 cells in 2 ml fresh Sl4 -m220 culture medium. 10. Count an aliquot of the cells (UNIT 1C.3) and dilute the Sl4 -m220 cell suspension to a concentration of 4 × 105 cells/ml. 11. Plate 0.5 ml from the cell suspension onto pregelatinized wells of 4-well culture dishes and incubate. 12. Next day, wash Sl4 -m220 cells with 0.5 ml CMF-PBS and replace with 0.5 ml PGC growth medium supplemented with both LIF (1000 U/ml) and FGF-2 (25 ng/ml). Reprogramming PGC to EG Cells

13. Use these plates for adding PGC suspension (see Basic Protocol).

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SEXING OF EG CELL LINES OR EMBRYOS BY PCR Established EG cell lines or early embryos (11.5 dpc) are sexed by PCR (Chuma and Nakatsuji, 2001). This technique is fast and reliable. Before isolation of genomic DNA from EG cells, somatic feeder cells have to be removed either by culturing the EG cells, for at least 2 passages, on gelatinized culture dishes in medium with LIF at twice the normal concentration (2000 U/ml) or by purifying EG cells from feeders using magnetic beads (see Support Protocol 1). Once feeder cells are not seen in EG cell cultures, genomic DNA is extracted and used for PCR analysis. For genotyping of embryos, the head is used as a source of genomic DNA. This technique is adopted from that previously published by Chuma and Nakatsuji (2001).

SUPPORT PROTOCOL 3

Materials 10-cm dish of EG cells without feeder cells PCR reagents including: 10 mM dNTP solution (10 mM each dNTP) 10× PCR buffer (containing 50 mM MgCl2 ; Qiagen, cat. no. 201203) 5 U/µl Taq DNA polymerase Autoclaved distilled water Primers: Ube1XA: TGGTCTGGACCCAAACGCTGTCCACA Ube1XB: GGCAGCAGCCATCACATAATCCAGATG Template genomic DNA DNA size ladder: 1-kb DNA ladder (Invitrogen) 2-µl thin-walled PCR tubes Thermal cycler (e.g., PTC-100, MJ Research) Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2000) 1. Isolate genomic DNA from one 10-cm culture dish of EG cells (Sambrook and Russell, 2001). Genomic DNA is isolated from EG cells grown on one gelatinized culture dish in medium containing a higher concentration of LIF (2400 U/ml) in order to remove feeder cells.

2. Set up a PCR reaction in 0.2-ml thin-walled tubes kept on ice:

15.5 µl distilled water 2.5 µl 10× PCR buffer (containing 15 mM MgCl2 ) 0.25 µl 50 mM MgCl2 1 µl 10 mM dNTPs 2.5 µl 10 pmol/µl Ube1XA primer 2.5 µl 10 pmol/µl Ube1XB primer 0.5 µl template DNA (genomic EG cells DNA diluted in water 1:10) 0.25 µl Taq DNA polymerase (5 U/µl) Total volume is 25 µl. When using a thermal cycler with a heated lid, do not use mineral oil.

3. Transfer the tubes to a PCR block and run the PCR program. 1 cycle: 35 cycles:

1 cycle 1 cycle

1 min 30 sec 30 sec 1 min 5 min indefinite

94◦ C 94◦ C 66◦ C 72◦ C 72◦ C 4◦ C

(denaturing) (denaturing) (annealing) (extension) (final extension) (hold).

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4. After the PCR amplification add 2 µl of loading buffer to 10 µl of the PCR products and run out on 2% agarose gel (Voytas, 2000) along with the 1-kb DNA marker. The size of PCR products are: male has two bands, 217 and 198 bp. Female has just 217 bp band.

Markers of PGCs and EG cells There are several markers used to identify both PGCs and EG cells. Nontissue specific alkaline phosphatase (AP) activity, the stage-specific embryonic antigen 1 (SSEA-1), germ cell nuclear antigen 1 (GCNA1), EMA-1, Oct-4, and nanog are used to identify PGCs or undifferentiated EG cells. These markers are downregulated in differentiated EG cells and in late PGCs (from 13.5 dpc onwards). A gene specifically expressed by PGCs is mouse vasa homolog (Mvh), which is expressed by 11.5 dpc PGCs onwards. Mvh antibody is commercially available from Abcam. SUPPORT PROTOCOL 4

NONTISSUE SPECIFIC ALKALINE PHOSPHATASE (AP) STAINING PGCs (from 8.5 to 12.5 dpc) and EG cells express a high level of tissue nonspecific alkaline phosphatase (AP; Fig. 1A.3.1B). Upon EG cell differentiation AP activity is downregulated. Here we present a protocol for performing AP staining on PGCs and EG cells using a commercially available kit.

Materials PGC from day 8.5 dpc embryos or EG cells grown on feeders PGC growth medium (see recipe) EG cell growth medium (see recipe) CMF-PBS AP staining solution (see recipe) 4-well culture dish Microscope 1. Culture 8.5 dpc PGCs in PGC growth medium supplemented with LIF (1000 U/ml) and FGF-2 for 1 to 5 days in a 4-well culture dish or grow EG cells on feeder cells in EG cell growth medium supplemented with LIF (1000 U/ml) for 2 to 3 days in a 4-well culture dish. 2. Aspirate the medium and rinse wells twice, each time with 0.5 ml of CMF-PBS. 3. Fix cells by air drying for 15 to 20 min at room temperature. If you want to preserve stained cells for more days, fix cells with 4% paraformaldehyde for 5 min at room temperature.

4. Make the AP staining solution. 5. Add 250 µl of the staining solution to each well of a 4-well dish. Keep in dark for 10 to 15 min to develop the red staining product. 6. Identify PGCs or EG cells (Fig. 1A.3.2C) by their dark red staining. SUPPORT PROTOCOL 5

IMMUNOFLUORESCENCE STAINING FOR SSEA-1 OR OCT-3/4 EG cells express many genes associated with pluripotency such as Oct-3/4, Sox-2, nanog, Esg-1, and SSEA-1. Upon EG cell differentiation, pluripotency genes are down regulated. Since Oct-3/4 and SSEA-1 antibodies are mouse antibodies, the staining is preformed separately.

Reprogramming PGC to EG Cells

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Materials CMF-PBS 4% (w/v) fixative solution (see recipe) PBS-TX solution (see recipe) SSEA 1 antibody (mouse, 1:1; Developmental Studies Hybridoma Bank, http://dshb.biology.uiowa.edu) Anti-Oct 3/4 (mouse, 1:250) from BD Transduction Laboratories Anti-mouse IgM-Alexa (red, 1:500; Molecular Probes) for SSEA-1 Anti-mouse IgG-Alexa (red, 1:500; Molecular Probes) for OCT-3/4 TOTO-3 solution (see recipe) Mounting medium for fluorescence (Vectashield; Vector) Lab-tek chambers (Nunc) for staining of cultured EG cells or PGCs or for freshly prepared cell suspensions use multiwell microscope glass slides (C.A. Hendley, cat. no. PH-136) precoated with poly-L-lysine to enhance the adherence of cells Humidified dark chamber Coverslips Fluorescence microscope Culture EG cells 1. Culture EG cells in Lab-tek chambers on MEFs. 2. Wash cells twice, each time with 0.25 ml CMF-PBS.

Fix EG cells 3. Remove the excess PBS and add 0.25 ml fixative solution for 15 min at room temperature. 4. Remove the fixative solution and wash wells twice, each time with 0.25 ml CMF-PBS for 5 min each. 5. Permeabilize and block cells with 0.25 ml PBS-TX for 20 min at room temperature.

Add primary and secondary antibody 6. Add either 150 µl SSEA-1 (1:1) or 150 µl Oct-3/4 (1:250) antibody diluted in PBS-TX. 7. Incubate overnight at 4◦ C in a humidified chamber. 8. Next day wash samples twice, each time with 0.25 ml CMF-PBS for 10 min each. 9. Add 200 µl of appropriate secondary antibody diluted in PBS-TX and incubate in a humidified dark chamber for 60 min at room temperature. 10. Wash twice, each time with 0.25 ml CMF-PBS for 10 min each. 11. Counterstain nuclei with 200 µl TOTO-3 solution for 15 min at room temperature in a humidified chamber in the dark. 12. Aspirate TOTO-3 solution and place a drop of fluorescent mounting medium on the slide and place a coverslip on top. Remove any excess mounting medium with paper tissue.

Examine samples 13. Examine samples under a fluorescence microscope with appropriate filters (Fig. 1A.3.3A, B), as soon as possible as the signal diminishes over time. Store slides at 4◦ C (short-term storage) or in a freezer at −20◦ C (for a few days) in the dark.

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Figure 1A.3.3 Characterization of pluripotent EG cells. Expression of SSEA-1 and Oct-3/4 was determined by immunofluorescence staining. EG cells express the cell surface antigen SSEA-1 (A, red), and Oct-3/4 (B, red). DNA is stained with TOTO-3 (blue).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Alkaline phosphatase (AP) staining solution Use the kit available from Sigma (cat. no. 86-R). Add 50 µl of sodium nitrite solution to 2.25 ml of autoclaved water and mix. Then add 50 µl of FRV-alkaline solution and 50 µl of naphthol AS-BI alkaline solution. Use the staining solution immediately.

Cell sorting medium 20 ml of cold DMEM (Invitrogen) supplemented with: 200 µl fetal bovine serum (FBS) 200 µl of 100× penicillin G + streptomycin (Invitrogen; 1× final) 200 µl of 100× L-glutamine (Invitrogen) Store up to 1 day at 4◦ C EG cell growth medium Add 6 ml of FBS to 32 ml of DMEM (Invitrogen, cat. no. 41965). Then add the following supplements:

400 µl of 200 mM L-glutamine 400 µl of 100 mM nonessential amino acids (NEAA;) 400 µl of 100× penicillin G + streptomycin (Invitrogen; 1× final) 400 µl of 100 mM sodium pyruvate 400 µl 2-mercaptoethanol solution (see recipe) 4 µl of 1000 U/ml LIF (see recipe) Store up to 2 weeks at 4◦ C Use FBS that has been cell culture tested.

Fixative solution, 4%

Reprogramming PGC to EG Cells

Working in a fume hood, dissolve 2 g of paraformaldehyde (PFA) in 35 ml water and warm up to 60◦ C in the oven. Then add 20 µl of 10% NaOH (to dissolve PFA) and put back into the oven. After 10 min, when PFA is dissolved, fill with water to 40 ml, cool to room temperature and add 5 ml of 10× PBS. Titrate to pH 7.4 with 1 N HCl. Fill to 50 ml with water. Filter using a 0.22-µm filter. Store up to 1 day at 4◦ C. Paraformaldehyde fixation preserves structure as well as retaining antigen recognition.

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FGF-2 stock solution Reconstitute 10 µg of FGF-2 (human recombinant; Invitrogen, cat. no. 13 256-029) in 1 ml of PBS. Store 60-µl aliquots up to 1 year at −20◦ C. FGF-2 is also known as basic fibroblast growth factor (bFGF).

Freezing solution Add 4 ml of DMSO (dimethyl sulfoxide, Sigma, D2650) to 6 ml of DMEM (Invitrogen) and mix. Then transfer into 30 ml of FBS and mix well. Keep freezing solution in −20◦ C freezer. Before use thaw, mix, and apply appropriate volumes for freezing the cells. Store the remaining freezing solution up to 1 year at −20◦ C. CAUTION: DMSO is a powerful solvent that can penetrate many synthetic and natural membranes, including skin and gloves. Therefore, potentially harmful substances, such as carcinogens, could be carried into the body through the skin or even gloves. For one 35-mm culture dish use 1 ml of freezing solution.

Gelatin, 0.1% (w/v) Dissolve 0.1 g of gelatin powder (swine skin type II, Sigma) in 100 ml of water (Invitrogen, cat. no. 15230), autoclave and store at room temperature.

Gelatinized petri dishes/wells Gelatinize dishes with 5 ml of 0.1% gelatin (see recipe) or 0.5 ml for wells in a 4-well plate for 30 min at room temperature and then remove the excess solution.

Leukemia Inhibitory Factor (LIF, ESGRO) Make 50-µl aliquots of LIF (107 U/1 ml; Chemicon International, cat. no. ESG1107) and store up to 1 year at 4◦ C. LIF is used at the concentration of 1000 to 1200 units in 1 ml of culture medium.

MEF medium 10% (w/v) FBS (Invitrogen) in DMEM Store up to 2 weeks at 4◦ C 2-Mercaptoethanol solution Add 7 µl of 2-mercaptoethanol (Sigma) to 10 ml of PBS. Sterilize using 0.22-µm filter. Store up to 1 week at 4◦ C.

Mitomycin C stock solution Make a stock solution of mitomycin C (Sigma, cat. no. M0503) by dissolving 2 mg in 1 ml of tissue culture–grade water by vortexing until dissolved. Then divide into 70-µl aliquots and store up to 6 months at −20◦ C.

PBS-TX solution Dissolve bovine serum albumin (BSA, Sigma, cat. no. A-7638) in PBS at a concentration 10 mg/ml. Add Triton X-100 to a final concentration of 0.1% (v/v). Store up to 1 week at 4◦ C. BSA is a blocking agent that reduces nonspecific antibody binding. Triton X-100 is a detergent used to permeabilize plasma and nuclear membranes. This solution is used for both blocking and permeabilization. Embryonic and Extraembryonic Stem Cells

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PGC growth medium Add 6 ml of FBS to 32 ml of DMEM (Invitrogen, cat. no. 41965). Then add the following supplements:

400 µl of 100× L-glutamine (Invitrogen, cat. no. 25030-024) 400 µl of 100× nonessential amino acids (NEAA; Invitrogen, 100×, cat. no. 11140) 400 µl of 100× penicillin G + streptomycin (Invitrogen, cat. no. 15140-122; 1× final) 400 µl of 100 mM sodium pyruvate (Sigma, cat. no. S8636) 400 µl 2-mercaptoethanol solution (see recipe) 4 µl of 1000 U/ml LIF (see recipe) 25 ng/ml of FGF-2 (see recipe) Store up to 2 weeks at 4◦ C Use FBS that has been cell culture tested.

Sl 4 m-220 medium 36 ml of DMEM (Invitrogen, cat. no. 41965) supplemented with: 4 ml FBS 400 µl of 100× L-glutamine (Invitrogen) 400 µl of 100× penicillin G + streptomycin (Invitrogen; 1× final) 400 µl of sodium pyruvate (Invitrogen) Store up to 2 weeks at 4◦ C TOTO-3 solution Add 1 µl of TOTO-3 (Molecular Probes, cat. no. T3604) to 500 µl of PBS. TOTO-3, a fluorescent DNA stain, is used to detect the nucleus in interphase cells and the chromosomes in mitotic cells.

COMMENTARY Background Information

Reprogramming PGC to EG Cells

PGCs are highly specialized cells that will eventually form sperm or eggs in the adult. Thus, they are also the only cell lineage that transmits both genetic and epigenetic information to the next generation. In the mouse, germ cell lineage is specified from the epiblast at 7.5 dpc. The specification and maintenance of germ cell lineage is critically dependent on the Blimp1/Prmt5 repressive complex (Ohinata et al., 2005; Ancelin et al., 2006). PGCs then migrate through the hindgut mesentery into the genital ridges, future gonads, which they reach by 10.5 to 11.5 dpc. PGCs during their development undergo epigenetic changes including erasure of histone H3 lysine 9 dimethylation (H3K9me2) and progressive upregulation of histone H3 lysine 27 trimethylation (H3K27me3; Seki et al., 2007). Upon PGC entry into genital ridges further epigenetic changes occur including reactivation of the inactive X chromosome in the female embryos and erasure of DNA methylation marks associated with imprinted genes and genomewide demethylation (Hajkova et al., 2002).

At the 12.5-dpc stage the sex of the embryo can be distinguished by morphology when the male genital ridge has stripes and the female genital ridge has spots (Chiquoine, 1954). At 13.5 dpc, PGCs enter meiotic prophase and mitotic arrest in both female and male, respectively. PGCs are not stem cells because they do not self-renew and do not make chimeras (Durcova-Hills et al., 2006) despite their expression of Oct-4, Sox-2, and nanog (Sch¨oler et al., 1990; Avilion et al., 2003; Yamaguchi et al., 2005). However, when PGCs are cultured in the presence of LIF, FGF-2, and SCF they can be induced to dedifferentiate into pluripotent stem cells, called embryonic germ (EG) cells, (Matsui et al., 1992; Resnick et al., 1992). It has been suggested that generation of EG cells in vitro and formation of germ cell tumors in vivo have many similarities (Donovan and de Miguel, 2003). The mechanisms underlying the molecular and epigenetic reprogramming are not well described. However, even the timing of the reprogramming was not known until recently when it was determined that cells

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coming from PGC formed colonies after only 10 days of culture are pluripotent stem cells because they made chimeras and were further propagated leading to derivation of EG cell lines (Durcova-Hills et al., 2006). Moreover exogenous FGF-2 was required only for the first 24 hr of PGC culture (Durcova-Hills et al., 2006) and the efficiency of making EG cell lines decreases with the gestational age of cultured PGCs (Durcova-Hills and McLaren, unpub. observ.). Several mutations are known to increase efficiency of generating EG cells including Pgct1 and Pten (Muller et al., 2000; Kimura et al., 2003). Established EG cell lines are propagated on mitotically inactive MEFs in only LIFsupplemented culture medium. EG cell lines have been derived from premigratory PGCs (8.0 to 8.5 dpc; Matsui et al., 1992; Resnick et al., 1992; Labosky et al., 1994), migrating PGCs (9.5 to 10.5 dpc; Durcova-Hills et al., 2001), or after their entry into the genital ridge (11.5 to 12.5 dpc; Tada et al., 1998; DurcovaHills et al., 2001) of different mouse strains, including 129Sv, C57BL/6, and strains of mixed background (Tada et al., 1998; Durcova-Hills et al., 2001). Recently it has been shown that pluripotent stem cell lines can be derived from neonatal testis; however, the growth factor requirements are different from that required for EG cells generation (Kanatsu-Shinohara et al., 2004). When EG cells are injected into blastocysts they contribute to germ line chimeras (Stewart et al., 1994) and upon omitting LIF from culture media they differentiate in vitro into all three embryonic germ layers including germ cell lineage (Resnick et al., 1992; Durcova-Hills et al., 2003). Mouse EG cells are similar to mouse ES cells. Both cell types express markers of pluripotency, form large multicellular colonies, and differentiate in vivo and in vitro into three germ layers. However, they differ in their DNA methylation status, where imprinted genes are hypomethylated in EG cells in contrast to ES cells (Tada et al., 1998; Durcova-Hills et al., 2001; Shovlin et al., 2008). EG cells as well as ES cells have the capacity to reprogram somatic cells into pluripotency after cell fusion (Tada et al., 1997, 2003). Recently EG cell lines have been derived in other species including human (Table 1A.3.1).

Critical Parameters All tissue culture is performed in a Class II Biological Hazard laminar-flow hood. Work surfaces are decontaminated before and after

each procedure with 70% ethanol. All reagents and media must be sterilized either by autoclaving or filtering through a 0.22-µm filter. Bottles and glass pipets are washed using a technique that avoids leaving residues of soaps or detergents, and are sterilized prior to use. Use batch-tested FBS, tested both for supporting the growth of derived pluripotent cell lines (ES cells or EG cells) and for the ability to support derivation of EGC-like colonies from cultured PGCs. Tested FBS should be aliquoted into 50-ml bottles and stored at −20◦ C. Thaw FBS only once to make medium and store any leftover serum at 4◦ C. Culture media containing all supplements should be discarded after 2 weeks or when the color turns purple, because of pH change. Keep separate bottles of medium for every researcher in the laboratory. Examine cultures and media daily for evidence of gross bacterial or fungal contamination.

Troubleshooting The undifferentiated EG cell colony usually has a round or oval shape with a distinct border around the colony in which individual cells are tightly compacted. Signs of differentiation can be observed by the changing morphology of colonies when they become flattened or when colonies adopt a more rounded shape and some free-floating embryoid bodies are formed. Markers of pluripotency are down-regulated during differentiation as assessed by AP, Oct-3/4, SSEA-1 immunofluorescence staining. When a culture starts to spontaneously differentiate it should be discarded. If the culture is very important, subculture only those colonies that have proper EG cell colony phenotype, using a mouth pipet (see the Basic Protocol). After 3 to 4 days, only EGC-like colonies should be observed in the culture. Expand the cells and freeze.

Anticipated Results The efficiency of EG cell line derivation declines with the gestational stage of embryos from which PGCs are isolated. PGCs isolated from an 8.5 dpc embryo are the most efficient for giving rise to EG cell lines. Approximately 4 to 6 EGC-like colonies are obtained from one PGCs-containing fragment isolated from an 8.5 dpc embryo which can individually be expanded and generate 6 EG cell lines (G. Durcova-Hills, unpub. observ.). Before starting to derive EG cell lines it is suggested to check PGC proliferation and formation of small and later larger PGC colonies (3 to 5 and later 10 to 20 cells) during the first 10 days of culture. Only a small subset of cultured PGCs

Embryonic and Extraembryonic Stem Cells

1A.3.17 Current Protocols in Stem Cell Biology

Supplement 5

Table 1A.3.1 Summary of EG Cell Lines Derived in Different Speciesa

Species PGC stage

Culture conditions

Chimeras

References

Mouse

8.5 dpc base of allantois 8.5 dpc 11.5 and 12.5 dpc 9.5 dpc

Sl4 cells, LIF+FGF-2+SCF →LIF STO cells, LIF+FGF-2+SCF →LIF Sl4 cells, LIF+FGF-2+SCF →LIF Sl4 cells, LIF+FGF-2+SCF →LIF

Germ line Germ line Germ line Somatic

Matsui et al. (1992) Resnick et al. (1992) Tada et al. (1998) Durcova-Hills et al. (2001)

Human

5-9 week GR 7-9 week GR

STO cells, LIF+FGF-2+forskolin→ LIF+FGF-2+forskolin STO cells, LIF+FGF-2+forskolin→ LIF+FGF-2+forskolin

ND ND

Shamblott et al. (1998) Turnpenny et al. (2003)

Pig

24-25 days GR 26-28 days GR 25-27 days GR 22-28 days GR

STO cells, LIF STO cells, LIF, FGF-2, SCF STO cells, LIF, FGF-2, SCF STO cells

Somatic Somatic Somatic

Shim et al. (1997) Tsung et al. (2003) Piedrahita et al. (1998) Rui et al. (2004)

Chicken Stage 28, 5.5 days of incubation Stage 28, 5.5 days of incubation

LIF+FGF-2+SCF+IL-11+IGF-1 LIF+FGF-2+SCF+IL-11+IGF-1

Somatic Germ line

Park and Han (2000) Park et al. (2003)

Goat

STO cells, LIF+SCF Goat embryonic fibroblasts

Somatic Somatic

K¨uhholzer et al. (2000) Jia et al. (2008)

BEFFCs, LIF, FGF-2, SCF

ND

Huang et al. (2007)

32 days GR 28-42 day of pregnancy

Buffalo 30-90 dpc GR

a Abbreviations: BEFFC, buffalo embryonic fibroblast feeder cells; GR, genital ridges; ND, not determined.

Reprogramming PGC to EG Cells

will give rise to EG cells. PGCs usually die off after 5 to 7 days of culture. When AP staining is used to identify PGCs we cultured PGCs cells in wells of a 4-well plate or in Lab-tek chambers. However, when SSEA-1 or Oct-4 is used to identify cultured PGCs by immunofluorescence, we culture PGCs in wells of a Labtek system. Cells used as feeders are negative for AP staining. PGCs form small colonies, which are enlarged in size in due course of the culture period.

eventually will give rise to an EG cell line. This can take up to 2 months.

Time Considerations

Sl 4 -m220 cells or STO cells Mitomycin C inactivation will require ∼2.5 hr. Sl4 -m220 cells can be obtained from Dr. Matsui from the Cell Resource Center for Biomedical Research, Institute of Development, Aging and Cancer, Tohoku University, Japan. STO cells can also be used for derivation of EG cell lines. STO cells are commercially available from European Collection of Cell Cultures (http://www.ecacc.org.uk).

EG cell line Generation of EG cell line is time consuming and labor intensive. Cultured PGC on feeders expressing SCF and in the presence of LIF and FGF-2 proliferate, and some of them form multicellular colonies resembling EG cell colonies after 8 to 10 days of culture. Subsequently, the individual colony comprising of 200 to 300 cells is further expanded and

Media Approximately 1 to 1.5 hr should be allowed for media preparation every week. Purification of PGCs using magnetic beads To isolate genital ridges from embryos and to make a single-cell suspension requires ∼2 hr depending on the number of embryos needed. The magnetic cell sorting will take an additional 2 hr.

1A.3.18 Supplement 5

Current Protocols in Stem Cell Biology

Literature Cited Ancelin, K., Lange, U.C., Hajkova, P., Schneider, R., Bannister, A.J., Kouzarides, T., and Surani, M.A. 2006. Blimp1 associates with Prmt5 and directs histone arginine methylation in mouse germ cells. Nat. Cell Biol. 6:623-630. Avilion, A.A., Nicolis, S.K., Pevny, L.H., Perez, L., Vivian, N., and Lovell-Badge, R. 2003. Multipotent cell lineages in early mouse development depend on Sox2 function. Genes Dev. 17:126140. Chiquoine, A.D. 1954. The identification, origin, and migration of the primordial germ cells in the mouse embryo. Anat. Rec. 118:135-146. Chuma, S. and Nakatsuji, N. 2001. Autonomous transition into meiosis of mouse fetal germ cells in vitro and its inhibition by gp130-mediated signaling. Dev. Biol. 229:468-479. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Donovan, P.J. and de Miguel, M.P. 2003. Turning germ cells into stem cells. Curr. Opin. Genet. Dev. 13: 463-471. Durcova-Hills, G., Ainscough, J. F.-X., and McLaren, A. 2001. Pluripotent stem cells derived from migrating primordial germ cells. Differentiation 68:220-226. Durcova-Hills, G., Wianny, F., Merriman, J., Zernicka-Goetz, M., and McLaren, A. 2003. Developmental fate of embryonic germ cells (EGCs), in vivo and in vitro. Differentiation 71:135-141. Durcova-Hills, G., Adams, I.R., Barton, S.C., Surani, M.A. and McLaren A. 2006. The role of exogenous FGF-2 on the reprogramming of primordial germ cells into pluripotent stem cells. Stem Cells 24:1441-1449. Hajkova, P., Erhardt, S., Lane, N., Haaf, T., Reik, W., Walter, J., and Surani, M.A. 2002. Epigenetic reprogramming in primordial germ cells. Mech. Dev. 117:15-23. Huang, B., Xie, T.-S., Shi, D.-S., Li, T., Wang, X.L., Mo, Y., Wang, Z.-Q., and Li, M.-M. 2007. Isolation and characterization of EG-like cells from Chinese swamp buffalo (Bubalus bubalis). Cell Biol. Inter. 31:1079-1088. Jia, W., Yang, W., Lei, A., Gao, Z., Yang, C., Hua, J., Huang, W., Ma, X., Wang, H., and Dou, Z. 2008. A caprine chimera produced by injection of embryonic germ cells into blastocyst. Theriogeneology 69:340-348. Kanatsu-Shinohara, M., Inoue, K., Lee, J., Yoshimoto, M., Ogonuki, N., Miki, H., Baba, S., Kato, T., Kazuki, K., Toyokuni, S., Toyoshima, M., Niwa, O., Oshimura, M., Heike, T., Nakahata, T., Ishino, F., Ogura, A., and Shinohara, T. 2004. Generation of pluripotent stem cells from neonatal mouse testis. Cell 119:1001-1012. Kimura, T., Suzuki, A., Fujita, Y., Yomogida, K., Lomeli, H., Asada, N., Ikeuchi, M., Nagy, A., Mak, T.W., and Nakano, T. 2003. Conditional loss of PTEN leads to testicular teratoma

and enhances embryonic germ cell production. Development 130:1691-1700. Kuhholzer, B., Baguisi, A., and Overstrom, E.M. 2000. Long-term culture and characterization of goat primordial germ cells. Theriogeneology 53:1071-1079. Labosky, P. A., Barlow, D.P., and Hogan, B.L.M. 1994. Mouse embryonic germ (EG) cell lines: Transmission through the germline and differences in the methylation imprint of insulin-like growth factor 2 receptor (Igf2r) gene compared with embryonic stem (ES) cell lines. Development 120:3197-3204. Matsui, Y., Zsebo, K., and Hogan, B.L.M. 1992. Derivation of pluripotential embryonic stem cells from murine primordial germ cells in culture. Cell 70:841-847. Muller, A.J., Teresky, A.K., and Levine, A.J. 2000. A male germ cell tumor-susceptibilitydetermining locus, pgct1, identified on murine chromosome 13. Proc. Natl. Acad. Sci U.S.A. 97:8421-8426. Ohinata, Y., Payer, B., O’Carroll, D., Ancelin, K., Ono, Y., Sano, M., Barton, S.C., Obukhanych, T., Nussenzweig, M., Tarakhovsky, A., Saitou, M., and Surani, M.A. 2005. Blimp1 is a critical determinant of the germ cell lineage in mice. Nature 439:207-213. Park, T.S. and Han, J.Y. 2000. Derivation and characterization of pluripotent embryonic germ cells in chicken. Mol. Reprod. Dev. 56:475482. Park, T.S., Hong, Y.H., Kwon, S.C., Lim, J.M., and Han, J.Y. 2003. Birth of germline chimeras by transfer of chicken embryonic germ (EG) cells into recipient embryos. Mol. Reprod. Dev. 65:389-395. Pesce, M. and De Felici, M. 1995. Purification of mouse primordial germ cells by MiniMACS magnetic separation system. Dev. Biol. 170:722725. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Piedrahita, J.A., Moore, K., Oetama, B., Lee, Ck., Scales, N., Ramsoondar, J., Bazer, F.W., and Ott, T. 1998. Generation of transgenic porcine chimeras using primordial germ cell-derived colonies. Biol. Reprod. 58:1321-1329. Resnick, J.L., Bixler, L.S., Cheng, L., and Donovan, P.J. 1992. Long-term proliferation of mouse primordial germ cells in culture. Nature 359:550551. Rui, R., Shim, H., Moyer, A.L., Anderson, D.L., Penedo, C.T., Rowe, J.D., BonDurant, R.H., and Anderson, G.B. 2004. Attempts to enhance production of porcine chimeras from embryonic germ cells and preimplantation embryos. Theriogeneology 61:1225-1235. Sambrook, J. and Russell, D.W. 2001. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.

Embryonic and Extraembryonic Stem Cells

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Supplement 5

Shamblott, M.J., Axelman, J., Wang, S., Bugg, E.M., Littlefield, J.W., Donovan, P.J., Blumenthal, P.D., Huggins, G.R., and Gearhart, J.D. 1998. Derivation of pluripotent stem cells from cultured human primordial germ cells. Proc. Natl. Acad. Sci. U.S.A. 95:1372613731. Sch¨oler, H. R., Dressler, G. R., Balling, R., Rohdewohld, H., and Gruss, P. 1990. A germline specific transcription factor mapping to the mouse t-complex. EMBO J. 9:2185-2195. Shim, H., Gutierrez-Adan, A., Chen, L-R., BonDurant, R.H., Behboodi, E., and Anderson, G.B. 1997. Isolation of pluripotent stem cells from cultured porcine primordial germ cells. Biol. Reprod. 57:1089-1095. Shovlin, T.C., Durcova-Hills, G., Surani, A., and McLaren, A. 2008. Heterogeneity in imprinted methylation patterns of pluripotent embryonic germ cells derived from pre-migratory mouse germ cells. Dev. Biol. 313:674-81. Seki, Y., Yamaji, M., Yabuta, Y., Sano, M., Shigeta, M., Matsui, Y., Saga, Y., Tachibana, M., Shinkai, Y., and Saitou, M. 2007. Cellular dynamics associated with the genome-wide epigenetic reprogramming in migrating primordial germ cells in mice. Development 134:2627-2638. Stewart, C.L., Gadi, I., and Bhatt, H. 1994. Stem cells from primordial germ cells can reenter the germ line. Dev. Biol. 161:626-628.

Tada, M., Tada, T., Lefebvre, L., Barton, S.C., and Surani, M.A. 1997. Embryonic germ cells induce epigenetic reprogramming of somatic nucleus in hybrid cells. EMBO J. 16:65106520. Tada, T., Tada, M., Hilton, K., Barton, S. C., Sado, T., Takagi, N., and Surani, M. A. 1998. Epigenotype switching of imprintable loci in embryonic germ cells. Dev. Genes Evol. 207:551-561. Tada, M., Morizane, A., Kimura, H., Kawasaki, H., Ainscough, J.F., Sasai, Y., Nakatsuji, N., and Tada, T. 2003. Pluripotency of reprogrammed somatic genomes in embryonic stem hybrid cells. Dev. Dyn. 227:504-510. Tsung, H.C., Du, Z.W., Rui, R., Li, X.L., Bao, L.P., Wu, J., Bao, S.M., and Yao, Z. 2003. The culture and establishment of embryonic germ (EG) cell lines from Chinese mini swine. Cell Research 13:195-202. Turnpenny, L., Brickwood, S., Spalluto, C.M., Piper, K., Cameron, I.T., Wilson, D.I., and Hanley, N.A. 2003. Derivation of human embryonic germ cells: An alternative source of pluripotent stem cells. Stem Cells 21:598-609. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Yamaguchi, S., Kimura, H., Tada, M., Nakatsuji, N., and Tada, T. 2005. Nanog expression in mouse germ cell development. Gene Express. Patterns 5:639-646.

Reprogramming PGC to EG Cells

1A.3.20 Supplement 5

Current Protocols in Stem Cell Biology

Derivation and Propagation of hESC Under a Therapeutic Environment

UNIT 1A.4

Kuldip S. Sidhu,1, 3 Sarah Walke,1 and Bernard E. Tuch1, 2 1

Diabetes Transplant Unit, The Prince of Wales Hospital and The University of New South Wales, New South Wales, Australia 2 New South Wales Stem Cell Network, New South Wales, Australia 3 Stem Cell Laboratory, Faculty of Medicine, School of Psychiatry, The University of New South Wales, New South Wales, Australia

ABSTRACT The pluripotent nature of human embryonic stem cells (hESC) makes them very attractive as a source of various cell types that could be used therapeutically in regenerative medicine. However, eliminating all sources of contamination, animal-derived or human cell–derived, during hESC derivation and propagation is necessary before hESC derivatives can be used clinically. Although there is continuing progress toward this goal, none of the methods to date to produce hESC lines under good manufacturing practices (GMP) has been published. The long-term success for GMP compliance depends critically on maintaining and implementing a stringent quality control system which is also dictated by the regulatory authorities in different countries. In this unit, an approach is described based upon the experience of this author and others towards achieving clinical-grade hESC lines systematically involving all the steps from start to finish under GMP environment. This unit provides a basic layout for GMP set up to achieve quality controls, a step-by-step guide to producing new hESC lines under defined conditions, and standard operating procedures used to achieve this outcome. Curr. Protoc. Stem Cell Biol. C 2008 by John Wiley & Sons, Inc. 6:1A.4.1-1A.4.31.
Keywords: hESC r GMP r standard operating procedures r quality control

INTRODUCTION This unit describes protocols for derivation of a human embryonic stem cell (hESC) line under a good manufacturing practice (GMP) environment that is largely based upon the author’s experience in producing a novel hESC line, called Endeavour-1, under culture conditions that are largely free of serum and animal products; these cells are propagated on a human feeder layer (Sidhu et al., 2008). Earlier attempts to replace murine fetal fibroblasts (MFF) with human source–derived fibroblasts as the feeder layer for hESC (Hovatta et al., 2003; Richards et al., 2003; Miyamoto et al., 2004; Park et al., 2004; Inzunza et al., 2005; Wang et al., 2005) all still used heterologous sera (e.g., FBS, KSR) and used immunosurgery to remove inner cell masses (ICM); hence they were not totally xeno-free systems. Similarly, feeder-free culture systems employed for isolation purposes used complex extracellular matrices—e.g., Matrigel, fibronectin, laminin— mostly derived from animal sources; hence these also were not xeno-free (Xu et al., 2001; Brimble et al., 2004; Rosler et al., 2004; Beattie et al., 2005). In these studies, either conditioned medium from mouse fibroblasts or growth factors (e.g., bFGF, TGFb, activin A, Nodal, Noggin, LIF, and PDGF) were used, and some degree of differentiation and chromosomal abnormalities were observed in hESC colonies in culture (Draper et al., 2004; Inzunza et al., 2004; Klimanskaya et al., 2005). Table 1A.4.1 summarizes recent advances towards a xeno-free culture of hESC. Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1A.4.1-1A.4.31 Published online July 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01a04s6 C 2008 John Wiley & Sons, Inc. Copyright


1A.4.1 Supplement 6

Table 1A.4.1 Summary of Recent Advances Toward Xeno-Free Culture of hESCsa

Cell lines Substrate

Key medium components

Longest time in culture

Characterizationb Mkr

Plur

Kary

Reference

Novel

STO cells

20% FBS

>30 passages

Yes

EB

Yes

Park et al. (2004)

Novel

Foreskin fibroblasts

20% FBS + LIF

9 months

Yes

Ter

Yes

Hovatta et al. (2003)

HES-3

FM fibroblasts

MEF-CM

>20 passages

Yes

Ter

Yes

Richards et al. (2002)

HES-4

FS fibroblasts

Novel

AFT epithelial cells

HES-3

AS fibroblasts

20% FBS

>30 passages

Yes

Ter

Yes

Richards et al. (2003)

HES-4

20% KSR

H1

Human marrow stromal cells

20% KSR

13 passages

Yes

EB

Yes

Cheng et al. (2003)

Novel

hES-df

20% KSR

44 passages

Yes

Ter, M

Yes

Stojkovic et al. (2005a)

Matrigel

hES-df-CM

14 passages

Yes

N/D

N/D

hES-df

20% KSR

18 passages

Yes

Ter, M

Yes

Matrigel

hES-df-CM

12 passages

Yes

N/D

N/D

H1

Matrigel

MEF-CM

6 months

Yes

EB

Yes

H7

Laminin

H1

Ter

H9

Xu et al. (2001) Rosler et al. (2004)

2 years

H14 MEF-CM

>24 passages

Yes

EB

Yes

Brimble et al. (2004)

Matrigel

HEF-TERT-CM

14 passages

Yes

EB

Yes

Xu et al. (2004)

Matrigel

40 ng/ml bFGF ± 15 passages other GFs

Yes

EB

Yes

Xu et al. (2005a)

BG01

Matrigel

BG02

Fibronectin

BG03 H1 H7 H9 H7

Derivation and Propagation of hESC Under a Therapeutic Environment

continued

1A.4.2 Supplement 6

Current Protocols in Stem Cell Biology

Table 1A.4.1 Summary of Recent Advances Toward Xeno-Free Culture of hESCsa, continued

Cell lines

Substrate

Key medium components

Longest time in culture

Characterizationb Mkr

H9 H1

Plur

Reference

Kary

Ter Matrigel

40 ng/ml bFGF + 33 passages 500 ng/ml Noggin

Yes

H9

EB

Yes

Xu et al. (2005b)

Ter

H14 H1

Matrigel

NIH/3T3-NogCM, 40 ng/ml bFGF + 500 ng/ml Noggin

I3

Human fibronectin

TGFβ1 ± LIF + >50 passages bFGF

7 passages

Yes

M

Yes

Wang et al. (2005)

Yes

EB

Yesc

Amit et al. (2004)

I6

Ter

H9 HSF6

Laminin

50 ng/ml activin >20 passages A, 50 ng/ml KGF, 10 mM NIC

Yes

Ter

Yes

Beattie et al. (2005)

H1

Matrigel

25 ng/ml activin A

Yes

EB

N/D

James et al. (2005)

H9

FBS

CDM + 10 ng/ml 10 passages activin + 12 ng/ml bFGF

Yes

EB

Yesc

Vallier et al. (2005)

H1

Matrigel Laminin

X-VIVO 10 + 80 >240 days ng/ml bFGF

Yes

Ter

Yes

Li et al. (2005)

H1

MEF-ECM

8% KSR + 8% 20 passages plasmanate + 16 ng/ml bFGF + 20 ng/ml LIF

Yes

EB

Yes

Klimanskaya et al. (2005)

BGN1 BGN2

H7 H9 4 others Novel

6 months continued

1A.4.3 Current Protocols in Stem Cell Biology

Supplement 6

Table 1A.4.1 Summary of Recent Advances Toward Xeno-Free Culture of hESCsa, continued

Cell lines

H1

Key medium components

Substrate

Longest time in culture

Characterizationb Mkr

Plur

Kary

Reference

Human serum

hES-df-CM

27 passages

Yes

M

Yes

Stojkovic et al. (2005b)

WA 15 & 16

Human Collagen IV, fibronectin, laminin and vitronectin/ feeder-free

Human serum 28 passages (albumin & transferrin)/DMEM/F12 + TGFβ + PA + GABA + LiCL + bFGF

Yes

M

No

Ludwig et al. (2006)

SA611

Human recombinant 20% Human gelatin/human serum/ feeder KO-DMEM + bFGF

>20 passages

Yes

M

Yes

Ellerstrom et al. (2006)

Endeavour- Human collagen 20% >40 passages 1 IV/novel serum-free KSR/KO-DMEM human feeder + bFGF

Yes

M

Yes

Sidhu et al. (2008)

Novel

Abbreviations: AFT, adult fallopian tube; AS, adult skin; bFGF, basic fibroblast growth factor; CDM, chemically defined medium (1:1 IMDM:F12 supplemented with insulin, transferrin, monothioglycerol and bovine serum albumin fraction V); FBS, fetal bovine serum; FM, fetal muscle; FS, fetal skin; GABA, gamma amino butyric acid; HEF-TERT-CM, conditioned medium from human ES cell-derived fibroblasts, stably transfected with TERT; hES-df, human ES cell-derived fibroblasts; hES-df-CM, human ES cell-derived fibroblast conditioned medium; KGF, keratinocyte growth factor; KSR, knockout serum replacement; LiCL, lithium chloride; LIF, leukemia inhibitory factor; MEF-CM, mouse embryonic fibroblast conditioned medium; MEF-ECM, extracellular matrix of MEFs; NIC, nicotinamide. PA, pipacholic acid. a Modified from Mallon et al., 2006 b Characterization key: Mkr, normal undifferentiated marker expression; Plur, pluripotency determined by embryoid body formation in vitro (EB), teratoma formation in vivo (Ter) or by monolayer differentiation in vitro (M); Kary, normal karyotype; N/D, not described. c Authors describe some abnormalities at late passage consistent with previous observations for cells grown on MEF feeders.

A number of hESC protocols are available for maintaining hESC lines—i.e., BresaGen hESC methods (http://stemcells.nih.gov/research/registry), ESI manual and other Singapore Protocols (http://www.stemcell.edu.sg/resources/methodsProtocols.php), Geron hESC methods (http://www.Geron.com/showpage.asp?code = prodstprot), Melton Laboratory hESC methods (http://mcb.harvard.edu/melton/HuES/), and WiCell hES Protocols (http://www.Wicell.org/forresearchers/index.jsp?catid = 12&subcatid = 20). The focus of this unit is to provide an outline for obtaining a GMPcompliant facility in the laboratory based on an Australian regulatory framework and to achieve the derivation of clinical-grade hESC lines. In principle this regulatory framework is not very different in other countries but there are some additional restrictions or the stringency in GMP compliance differs. URLs for representative authorities include: Australia, http://www.tga.gov.au/docs/html/gmpcodau.htm; Canada, http://www.hc-sc.gc.ca/dhp-mps/compli-conform/gmp-bpf/docs/index e.html; Europe, http://ec.europa.eu/enterprise/pharmaceuticals/eudralex/homev4.htm and USA, http://www.cgmp.com/howGmpsChange.htm). The appropriate authority should be consulted when setting up a laboratory for deriving hESC lines.

STRATEGIC PLANNING Derivation and Propagation of hESC Under a Therapeutic Environment

Designing a GMP-Compliant Facility for Production of hESC Lines A setup for a small-to-medium size academic or biotech laboratory is described; largescale manufacturing facilities for therapeutic purposes may require a different regulatory

1A.4.4 Supplement 6

Current Protocols in Stem Cell Biology

Figure 1A.4.1 A floor plan for two clean rooms and adjacent storage space in DTU Prince of Wales Hospital Australia (courtesy of Kuet Li and Sarah Walke).

framework. Bear in mind that hESCs are not any different from other cell types when setting up GMP facility. Figure 1A.4.1 gives a floor plan for two proposed clean rooms with adjacent storage facilities at the Diabetes Transplant Unit (DTU), Prince of Wales Hospital, Australia. To meet the regulatory requirements and GMP compliance, the clean rooms are generally designed and fabricated by professionals. Professional servicing is also available for maintaining the climate control, including environmental control (suspended particles, etc.), in clean rooms before the commencement of work. Changing from one cell type to another one in the same clean room is allowed in Australia after professional clean up (level of suspended particles) of the room, but this may be different in other countries. The long-term success for GMP compliance depends critically on maintaining and implementing a stringent quality control system which is also dictated by the regulatory authority in the country.

Standard Operating Procedures Apart from standard tissue culture practices, derivation and propagation of hESC require some specialized and standardized handling and culturing techniques (see Support Protocols 1 to 10). The procedures described in this unit, based on the experience of the author and others, are reliable and reproducible for obtaining meaningful experimental outcomes. These procedures also need reviewing and upgrading periodically, keeping in mind the new advances made in this field.

Embryonic and Extraembryonic Stem Cells

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Quality Control (QC) of Cell Type Produced, i.e., hESC Although hESC lines maintain a baseline for expression of stem cell surface markers and other characteristics, subtle differences in marker expressions between lines are observed if the cells are cultured over an extended period of time. The basic set of tests recommended for QC purposes are routine karyotyping, RT-PCR analysis of pluripotency markers (Nanog/OCT4), and differentiation markers for ectoderm, mesoderm, and endoderm—i.e., nestin, brachyury, α-fetoprotein, respectively—immunocytochemical analysis for stem cell surface markers (SSEA3/4, TRA-1-61, TRA-1-80), and alkaline phosphatase staining, demonstrating in vivo pluripotency by teratoma formation after injecting under the kidney capsule of SCID mice. In addition, hESC lines should also be routinely tested for mycoplasma, fungal, and bacterial contaminations. Maintaining Stocks, Cell Banking, and Distribution Maintaining quality control (QC) stocks of hESC lines in a well-structured cell bank with a well-defined database of labeled (identity, passage number, date) samples is an essential component of a good cell facility. Early passage hESC lines should be archived as mother stocks followed by essential and critical master stock, working stock, and the stock for experiments for regular use at different levels or tiers of liquid nitrogen storage tank. Liquid nitrogen tanks can be tucked away in the premises of a GMP facility, but a provision must be made to refill the tanks from outside the GMP premises (see Fig. 1A.4.1). Both slow and rapid freezing protocols (Support Protocols 5 and 6) can be used to cryopreserve hESC lines, with better post-thaw recovery reported with rapid freezing method. Work/Time Flow for Derivation of hESC Lines Production of hESC lines usually involves two separate institutions—infertility clinics for supply of eggs or inner cell masses (ICM) and a research laboratory for derivation of hESC lines from ICM—and these institutions are usually located separately. This situation makes it a bit difficult for GMP compliance on the final product unless work and time flow are properly coordinated. Figure 1A.4.2 gives a sample work/time flow in making hESC lines at DTU that meets GMP compliance on the final product as an

Derivation and Propagation of hESC Under a Therapeutic Environment Figure 1A.4.2

Work/time flow in making clinical-grade hESC lines.

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example. The transport of material between two institutions for derivation of hESC lines as indicated must be under proper climate control in portable and sealable CO2 incubator for GMP compliance. A sample weekly schedule for coordinating hESC and human fetal fibroblast (HFF) maintenance is described in Support Protocol 1. NOTE: All procedures should be performed under sterile conditions. All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All studies with human subjects must be approved by the Institutional Review Board (IRB), which must adhere to the Office for the Protection from Research Risk (OPRR) guidelines or other applicable governmental regulations for using human subjects. All material must be obtained with informed consent of the donor.

DERIVATION OF A NEW hESC LINE FROM HUMAN BLASTOCYSTS The derivation of new hESC lines from embryos that are in excess of a couple’s reproductive need is permissive, with informed consent, under legislation with a license in Australia as in many other countries. This procedure also requires necessary institutional ethics approval. Bear in mind that applying for such a license in collaboration with infertility clinics may be a very time consuming effort and must be carefully planned ahead so that milestones (obtaining license and producing hESC lines) are achieved and the project can go forward.

BASIC PROTOCOL

Materials Frozen human embryos, preferably blastocyst stage Quinn’s Advantage Cleavage and Blastocyst medium (SAGE BioPharma) supplemented with 5% human serum albumin (hSA; Sage Biopharma) Oil for tissue culture (SAGE BioPharma) SR medium plus bFGF (see recipe) Mitotically inactivated (γ-irradiated) human fetal fibroblasts (HFF) as feeder layer (see Support Protocols 7 and 8) 4- or 6-well culture plates (Greiner bio-one, GmbH, Germany) Inverted microscope (example, Leica DM-IRB) with CCD camera and software to manipulate images, and with laser ablation system (e.g., XYClones; Hamilton Thorne Biosciences) Portable CO2 incubator (LEC Instruments, http://www.lecinstruments.com/incubator.htm; see Fig. 1A.4.3) Dissection and biopsy pipets (e.g., Cook IVF) Water-Jacketed CO2 incubator (e.g., Gelaire, Sydney) Nalgene Cryofreezing Containers (Fisher Scientific, Nalgene cat. no. 5100-001) Microchisel, 10× (e.g., Eppendorf) or insulin syringe with 23-G needle Biological safety cabinet (BSC) with a provision to keep a microscope inside for performing hESC sub culturing Pipettor with 100-µl tip Additional reagents and equipment for preparing HFF feeder cells (Support Protocol 8) Thaw and culture embryos 1. Classify the embryos based on the appearance of the ICM at the blastocyst stage. Embryos in the infertility clinics are generally frozen at different stages of development (i.e., from two pronuclear to blastocyst stage) and they vary considerably in

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Figure 1A.4.3

A portable CO2 incubator.

Figure 1A.4.4 Diagrammatic representation of embryos category (A through E) based on appearance of inner cell mass.

quality. Assessment of embryo quality by an embryologist in the infertility clinic is helpful. Derivation and Propagation of hESC Under a Therapeutic Environment

Based on the appearance of ICM, embryos can be classified into five broad categories, A through E as represented in Figure 1A.4.4. Briefly, category A has a compact mass of cells indistinguishable from each other; category B, cells are not compact but loosely adhere together; category C, few cells difficult to distinguish from trophectoderm; category D, a few degenerative cells; and category E, no ICM visible.

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Figure 1A.4.5 Derivation of Endeavour-1 on serum-free HFFs used as feeder layer. (A) Serumfree HFFs as monolayer. (B) Normal growth of three hES colonies on serum-free HFFs. (C) Localization of alkaline phosphatase (marker of pluripotency) in a colony of Endeavour-1. (D) A normal-looking human embryo after thawing. (E) Hatched blastocyst with visible ICM (higher magnification in inset) (F) A nascent colony of Endeavour-1. (G) A fully grown colony of Endeavour1. (H) A normal-looking colony of Endeavour-1 after first passage. (I) Normal-looking colonies of Endeavour-1 at passage 9 (from Sidhu et al., 2008).

2. Thaw embryos rapidly in 4-well culture dishes, preferably by an embryologist in the infertility clinic, and culture in 20 µl Quinn’s Advantage Cleavage and Blastocyst medium supplemented with 5% hSA under oil until they develop into blastocysts (Fig. 1A.4.5E). 3. Culture the blastocysts overnight for expansion and hatching. Alternately hatching could be assisted by a laser attached to the microscope.

Dissect ICM and co-culture on feeder cells 4. Prepare HFF feeder cells in a 6-well plate with 2 ml of SR medium plus bFGF/well (see Support Protocol 8) a day before the start of ICM isolation from embryos. Transport the feeder cell plates to the infertility clinic in a portable 5% CO2 incubator (Fig. 1A.4.3). It is important to remember that for GMP compliance, HFF feeder layers are prepared in human-derived collagen IV-coated plates as apposed to animal-derived gelatin-coated plates.

5. Isolate the ICM. In a good quality hatched-blastocyst (category A), ICM can be visualized under the phase contrast microscope (Fig. 1A.4.5E).

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Figure 1A.4.6 A schematic for laser dissection of ICM from embryo. The XYClone ablation system shown is mounted on to the inverted microscope (not shown).

Various procedures are used to dissect out ICM—i.e., immunodissection using antibodies (Kim et al., 2005), physical dissection (see Rajan et al., 2007), laser dissection (Sidhu et al., 2008), or using whole embryo culture. Immunodissection has the limitation of introducing animal-derived products into the culture system and thus compromises GMP compliance. The authors have successfully used laser dissection of ICM from hatched blastocysts for generating a new hESC line, Endeavour-1 (Sidhu et al., 2008) as it eliminated the use of animal products, such as antibodies for immunosurgery. Briefly, the authors have used the XYClone system that incorporates a laser within 40× objective of a microscope and delivers a highly focused laser beam (Class 1, 1480 nm) to the targeted area resulting in precise ablation of desired cells. The laser beam (red with effective beam area shown in pink color) has cushion (shown as yellow color) around it that protects against damage or trauma to the desired area (see Fig. 1A.4.6). It allows a very precise separation of ICM from trophoblast, maintaining the intactness of ICM. The ICMs are dissected from hatched blastocysts culture in Quinn’s Advantage Cleavage and Blastocyst medium supplemented with 5% hSA in a 4-well plate and using an XYClone Class I laser (HD Scientific Suppliers www.hdscientific.com.au) guided by phase contrast microscope. The flow of work/time is as described in Figure 1A.4.2. Each time 5 to 8 embryos could be processed simultaneously to obtain hESC lines.

6. Transfer the dissected out ICMs individually using a biopsy pipet (or pipettor with a wide-mouth tip) into each well of 6-well HFF feeder plate. Incubate until the ICMs attach. Attachment of ICM to feeder may take 2 to 3 days, culture is transferred to the research laboratory only after attachment of the ICM to the feeder layer. If ICM does not attach to feeder within 3 to 4 days, it is considered dead and discarded.

7. The following day, transfer the plate with ICMs that have attached to the feeder layer to the clean rooms of the research laboratory in a portable water-jacketed CO2 incubator (Fig. 1A.4.3) with a maximum total travel time of 30 to 45 min. Derivation and Propagation of hESC Under a Therapeutic Environment

Many companies offer a portable CO2 incubator; we use a local made inexpensive CO2 incubator that can house a 6-well plate and can maintain 5% CO2 and 37◦ C.

8. Daily, replace half of the medium in each well with fresh SR plus bFGF 4 ng/ml. Monitor growth of the ICM carefully over the next 10 to 14 days (see Fig. 1A.4.4G,H).

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Figure 1A.4.7

Manual dissection of hESC colony into smaller pieces using a microchisel.

9. When the outgrowth has formed a full-sized hESC colony i.e., 300 to 400 µm, dissect the hESC colony physically into 8 to 12 pieces by using microchisel or a sterile insulin syringe with a 23-G needle. Carry out the dissection under an inverted microscope in a biological safety cabinet (BSC), class II. Briefly, two to three vertical and two to three horizontal cuts are made first before making a peripheral cut along the rim of hESC colony (Fig. 1A.4.7).

10. Gently lift the 6 to 8 pieces of hESC colony and transfer them using a pipettor with a 100-µl tip to a fresh 6-well HFF feeder plate. 11. Continue manual dissection of hESC colonies until 40 to 50 moderate-sized (200 to 300 µm) colonies/well are obtained (usually in 5 to 6 passages). Each fragment gives rise to a colony and these colonies are all derived from one ICM.

12. Use some of these colonies for cryopreservation at this stage (see Support Protocol 5 or 6).

Establish a new hESC line Establishing a new hESC line may take several months as a stable line should survive repeated freeze/thaw cycles with good post-thaw recovery and maintain karyotype stability and pluripotency in culture. Generally it takes ∼10 to 12 passages before a stable new cell line is obtained. 13. Assess the colonies during establishment of new hESC lines: a. Recovery after cryopreservation using both slow and fast freezing (vitrification; see Support Protocols 5 and 6). About 50% to 75% recovery from fast freezing (fast freezing gives better post-thaw recovery) indicates a stable new line. Recovery after 5 to 7 freezing thawing cycles is a good indication of a stable new line.

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Figure 1A.4.8 Histological demonstrations of various tissues formed in the teratomas by Endeavour-1 after injecting under the kidney capsule of SCID mice and its karyotyping analysis. (A) neuroectoderm (arrow, ectoderm). (B) gut-like structures (arrow, endoderm). (C) cartilage-like structure (arrow, mesoderm). (D) Karyotype, 46XX (from Sidhu et al., 2008).

b. Karyotype stability: Karyotyping should be carried out after every 10th passage initially for the first 20 passages and then after 20 to 30 passages. A stable hESC line should maintain karyotype for an extended period (Fig. 1A.4.8D)

c. Optimal expression of stem cell surface markers: i.e., SSEA3/4 and low or no expression for SSEA1 (Fig. 1A.4.9), TRA-1-61, TRA-1-80 by immunocytochemistry (>95% cells should be positive), and by FACS analysis (>50% to 60% cells should be bright SSEA3/4 positive cells; Sidhu and Tuch, 2006). Immunofluoresence staining can be done according to the procedure described by Chemicon (http://www.chemicon.com). Cells can also be analyzed for alkaline phosphatase staining using the Dako (K 0624) kit for this purpose and following the instructions included with the kit (http://www.dako.com.au). The majority of cells (>99%) should be positive for alkaline phosphatase staining.

d. Gene (RT-PCR) and protein (immunofluorescence) expressions for pluripotent markers, i.e., Nanog/OCT4 should be assessed (Fig. 1A.4.10). e. Demonstration of pluripotency in vitro by embryoid body formation (Fig. 1A.4.10) and after differentiation analysis of gene expressions by RT-PCR of lineage markers, i.e., nestin (ectoderm), brachyury (mesoderm), and α-fetoprotein (endoderm). f. Similarly the demonstration of pluripotency in vivo by formation of teratomas after injecting hESC under the kidney capsule of SCID mice (Fig. 1A.4.8A-C). The teratomas should contain tissue derived from ectoderm, mesoderm, and endoderm.

g. If more than one line is maintained in the facility, HLA (human leukocyte antigen) typing and genomic fingerprint are also recommended. Derivation and Propagation of hESC Under a Therapeutic Environment

These assays are available as off-the-shelf kits or in any forensic/pathology laboratory.

h. Before distribution of these lines, cultures must be tested for microbial contamination from mycoplasma, fungi, and bacteria.

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Figure 1A.4.9 (A) Upper panel, Immunolocalization of stem cell surface markers. From left, OCT4, SSEA3, SSEA4, TRA-1-81, TRA-1-60 in clone hES 3.1. A similar expression of these surface markers was observed in other clones. Lower panel, FACS-sorted TRA-1-60 positive bright hESC. From left, hESC 3, Clone 3.1, Clone 3.2, Clone 3.3 (Magnification 400 ×; from Sidhu and Tuch, 2006). (B) FACS analysis of SSE1/4 expression in hESC. Panel B reprinted from The International Journal of Biochemistry and Cell Biology, volume 38, Mallon, B.S., Park, K.Y., Chen, K.G., Hamilton, R.S., and McKay, R.D., Toward xeno-free culture of human embryonic stem cells, pages 1063 to 1075, copyright 2006, with permission from Elsevier.

Figure 1A.4.10 Characterization of Endeavour-1 and its clonal lines, E1C1, E1C2, E1C3, and E1C4. Upper panel, RT-PCR expression of genes for pluripotency (Nanog) and lower panel, EBs (arrows) formed from E1 in suspension culture and their differentiation to different cell lineages (arrows). Similar cell types were also obtained after differentiation of clonal lines (adapted from Sidhu et al., 2008).

hESC WEEKLY CULTURE SCHEDULE This protocol is an example of a weekly schedule for maintaining hESCs. The tasks should be adapted to the schedule for the individual laboratory.

SUPPORT PROTOCOL 1

Materials HFF, cryopreserved (Support Protocol 9) hESC cultures (Basic Protocol) SR medium for both hESCs and HFF (see recipe) Collagen IV-coated 75-cm2 flasks (see recipe) Collagen IV-coated 6-well plates (see recipe) γ irradiator

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Friday 1. Thaw and plate cryopreserved HFF at passage 5 (p5) in collagen IV-coated 75-cm2 flasks for Monday. HFF at p 6 to 8 could also be used.

2. During the afternoon, change medium on all dishes containing hESCs using SR medium. Record colony morphology, number of fragments attached, and differentiation (appearance of cobble stone morphology in colonies) if any, on the data sheets. 3. Equilibrate required volume of SR medium for weekend medium change At a time point 24 hr prior to its use, SR medium should be equilibrated at 37◦ C, 5%CO2 . Under sterile conditions aliquot the volume required to change medium on all plates from prepared SR medium stock (maintained at 4◦ C) to a sterile 25-cm2 tissue culture flask and place in incubator. Equilibration of SR medium will lessen the effectiveness of the penicillin/streptomycin in the medium and increase the risk of contamination of the hESC cultures. Therefore totally aseptic conditions must be employed when changing medium.

Saturday or Sunday 4. Change medium on HFF plates set up on Friday with 2 ml/well of SR medium 5. Change medium on all 6-well plates containing hESC using 2 ml/well with equilibrated SR medium. Record colony morphology. 6. Equilibrate required volume of SR medium for Monday medium change.

Monday 7. Collagen IV–coat 6-well culture plates. 8. Harvest HFF set up on previous Friday (see Support Protocol 8). 9. Seed HFF into 6-well culture plates at 1.5 × 105 cell/ml using 2 ml/well SR medium 10. Change medium on all dishes containing hESC using equilibrated SR medium. Record colony morphology. 11. Equilibrate required volume of SR medium for Tuesday medium change. Be sure to include medium for 6-well culture plates set up today.

Tuesday 12. Check 6-well HFF culture plates set up Monday for any contamination. 13. γ irradiate 6-well HFF culture plates, 45 Gy 5 to 6 min 14. Change medium on all dishes containing hESC using 2 ml/well equilibrated SR medium. Record colony morphology. 15. Make up fresh SR medium if necessary. 16. Equilibrate required volume of SR medium for Wednesday medium change. Be sure to include SR medium for the new 6-well culture plates.

Wednesday 17. Change medium on all new feeder layer plates using 2 ml SR medium per well. 18. Change medium on all hESC cultures. Record colony morphology. Derivation and Propagation of hESC Under a Therapeutic Environment

19. Equilibrate required volume of SR medium for Thursday medium change.

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Thursday 20. In the morning, change medium on all new feeder plates using 2 ml/well equilibrated SR medium per well. Incubate the plates a minimum of 3 hr before transferring new colony fragments onto feeder layers. 21. Observe hESC plated on Thursday of last week and decide which to transfer to new 6-well HFF culture plates. 22. In the afternoon, divide and transfer hESC colonies to new HFF feeder plates. Initially mechanical passaging is recommended for 5 to 6 passages followed by enzymatic passaging (Support Protocols 3 and 4). 23. Record number of fragments transferred, collected for RNA, frozen down, or used for other experimental procedures on the data sheets. 24. On alternate weeks, perform division of hESC colonies for fast freezing (Support Protocol 6). 25. Equilibrate required volume of SR medium for Friday medium change.

DETERMINING VIABILITY OF hESC BY CARBOXYFLUORESCEIN DIACETATE (CFDA) AND PROPIDIUM IODIDE (PI)

SUPPORT PROTOCOL 2

The purpose of this protocol is to assess viability of hESCs during propagation using a vital dye, CFDA, that stains viable cells green and PI that stains nonviable cells red.

Materials Carboxyfluorescein diacetate (CFDA) DMSO Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen) Propidium iodide (PI) hESC removed from the culture dish (Basic Protocol) and placed in a microcentrifuge tube 1-ml pipettor 20- to 200-µl micropipettor Hemacytometer Fluorescent microscope equipped with UV filters 1. Prepare 10 mM 6-CFDA in DMSO (4.6 mg 6-CFDA/ml DMSO). Store this stock solution at 4◦ C. Before use dilute the stock solution 1:100 in PBS for use in this protocol. 2. Prepare PI at a concentration of 100 µg/ml in PBS by adding 1 mg of powdered PI to 10 ml of PBS and store this stock solution at 4◦ C. 3. Wash hESC twice by resuspending them in 0.5 ml PBS, microcentrifuge 3 min at 500 to 600 × g, room temperature. Carefully remove the supernatant with a 1-ml pipettor. 4. Add 250 µl of 6-CFDA to the hESC and incubate for 30 min in a 37◦ C incubator. 5. Wash with PBS by adding 0.5 ml PBS and microcentrifuge 3 min at 500 to 600 × g, room temperature. Carefully remove the supernatant with a 1-ml pipettor. Repeat. 6. Resuspend the cells in 200 µl of PBS. Embryonic and Extraembryonic Stem Cells

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7. Add 10 µl of 100 µg/ml PI and incubate 5 to 10 min at room temperature. Then place on ice. 8. Pipet 20 µl of the cell suspension, using a 20- to 200-µl micropipettor, under the coverslip on a hemacytometer. Visualize under the fluorescent microscope equipped with UV filters. 9. Estimate the percentage of hESCs which are viable (green fluorescence) and nonviable (red fluorescence). SUPPORT PROTOCOL 3

PASSAGING INTACT hESC COLONIES BY COLLAGENASE/DISPASE TREATMENT This procedure describes how to obtain intact undifferentiated hESC colonies required for EB formation, freezing, subculturing, lineage specification, and for producing teratomas to determine pluripotency in these cells. This procedure can also be used to obtain single-cell preparations from hESC colonies.

Materials hESCs cultured on HFF feeders in 6-well plates (Basic Protocol) Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen), prewarmed 1 mg/ml collagenase (Invitrogen) in PBS, sterilized with a 0.22-µm syringe filter and prewarmed 0.5 mg/ml dispase (Invitrogen, cat. no. 17105-041) in PBS, sterilized with a 0.22-µm syringe filter and prewarmed SR medium (see recipe), prewarmed to 37◦ C Trypsin/EDTA (Invitrogen, cat. no. 25300-054) or TrypLE Select (Invitrogen, cat. no. 12563-011), prewarmed Microscope Plastic loop (Lazy-L-Spreader; Cole-Parmer Instrument) 15-ml tube Biological safety cabinet (BSC) Class II hood 1. Aspirate medium from hESC cultures in 6-well plates and wash each well with 1 ml D-PBS twice. 2. Aspirate PBS. 3. Add 1 ml collagenase/well and incubate at 37◦ C in CO2 incubator. 4. Observe the plates under a microscope at 4× magnification, and when most of the hES colonies are sufficiently rounded up (∼10 min), proceed to the next step, otherwise keep incubating and checking every 10 min up to a maximum 30 min (7 to 10 min is optimum). If some differentiated hESCs colonies are present, first remove them (pick to loose, PTL) by dissecting out or simply by scratching using a plastic loop under the microscope and replacing PBS with fresh PBS. If only a small number of undifferentiated colonies is present, these can be picked (pick to keep, PTK) by dissection or by scratching under the microscope.

Derivation and Propagation of hESC Under a Therapeutic Environment

5. To completely lift the colonies it may be necessary to add 1 ml/well dispase for 5 min without removing collagenase. This step is not always necessary.

6. Remove collagenase and wash wells gently with 1 ml PBS.

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7. Add 1 ml fresh PBS/well and lift colonies by scratching gently with a plastic loop. Any colonies that are not removed the first time can be lifted again the same way by using another 1 ml of PBS

8. Transfer colonies to a 15-ml tube and let the colonies settle to the bottom of tube (5 min) in a BSC Class II hood. 9. Carefully aspirate PBS and replace it with 1 ml SR medium. These hESC colonies can be used for EB formation or for subculturing or for inducing teratomas in SCID mice.

Prepare a single-cell suspension 10. Treat ∼50 to 75 intact hESC colonies (picked up as above) with 100 µl of 0.05% trypsin or TrypLE Select for 7 min at 37◦ C. Make a single-cell suspension by frequently triturating with a pipet during the digestion. Neutralize trypsin at the end of the incubation by adding 1 ml SR medium. These single-cell suspensions are used for FACS analysis of SSEA3/4 positive cells.

SUBCULTURING hESC COLONIES BY TrypLE SELECT TREATMENT This procedure is used to scale up propagation of hESC by efficiently subculturing using TrypLE Select. The enzyme is a recombinant enzyme derived from a bacterial source and thus eliminates the source of animal-derived products in the cultures.

SUPPORT PROTOCOL 4

Materials hESCs cultured on HFF feeders in 6-well plates (Basic Protocol) Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen), prewarmed at 37◦ C TrypLE Select, prewarmed SR medium (see recipe), prewarmed Plastic loop (Lazy-L-Spreader, Cole-Parmer Instrument) 15-ml Falcon tube 1. Aspirate medium from 6-well plates and wash each well with 1 ml PBS twice. 2. Aspirate PBS. 3. Add 0.3 ml TrypLE Select per well. 4. Observe under microscope at 4× magnification, and when HFF layer is sufficiently rounded up (within 2 min), add 2 ml SR medium and gently pipet up and down to wash the wells until HFF layer is completely detached lifting hESC colonies. Lift colonies by scratching gently with a plastic loop. The carried-over HFF cells being irradiated will die subsequently.

5. Transfer the contents of each well to 15-ml Falcon tube and wash the wells with additional 1 ml SR medium. Pool the contents and make up the required volume for splitting the cells. A total of 12 ml is required for a six-well plate (2 ml/well). Normally, the split ratio is 1:6.

6. Triturate 5 to 7 times and aliquot into fresh 6-well plate containing HFF. Use a circular motion of pipet to evenly distribute hESC colonies in the well. Incubate until the next passage. Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 5

SLOW FREEZING hESC COLONIES/CLUMPS This method is used to cryopreserve hESC clumps at low passage number for later propagation and lineage specification studies. Vitrification can be used for a similar purpose but slow freezing is more conveniently performed and hence preferred.

Materials hESC colonies harvested by collagenase/dispase (Support Protocol 3) 30% SR medium [9 ml of 20% SR medium +1 ml KOSR (Invitrogen, cat. no. 10828-028)] Cryopreservation medium II: 6 ml Knockout DMEM (Invitrogen), 2 ml of 30% SR medium, 2 ml DMSO, sterile filtered using a 0.22-µm syringe filter HFF feeder plates (Support Protocol 8) 15-ml tube Cryovials (Greiner Bio-One, cat. no. 122263) Nalgene Cryofreezing Containers (Fisher Scientific, Nalgene 5100-001) −80◦ C freezer Cryoboxes (Crown Scientific) Liquid nitrogen tank 37◦ C water bath Freezing hESC colonies 1. Pick up colonies from each well released by collagenase/dispase treatment (Support Protocol 3). 2. Let the colonies settle at the bottom of a 15-ml tube (5 to 6 min) and remove as much of the supernatant as possible. This helps in removing HFFs.

3. Resuspend hESC colonies (50 to 75 colonies) from each well of the 6-well plate in 2 ml of 30% SR medium (9 ml of 20% SR medium + 1 ml KOSR) and gently break colonies to smaller but not too small pieces with a pipet. Four to six pieces per hESC colony, i.e., 25- to 30-µm colony pieces are used.

4. Dropwise add equivalent volume (2 ml) of cryopreservation medium. 5. Mix and transfer 1 ml each to labeled cryovials. Transfer vials into Nalgene container for overnight storage at −80◦ C. 6. The next day, transfer the vials to cryoboxes and store in liquid nitrogen tanks for long-term storage.

Thawing hESC colonies 7. Transfer cryovial from liquid nitrogen directly into a water bath at 37◦ C and thaw the contents as quickly as possible by shaking. 8. Transfer thawed contents into one well of 6-well plate containing 2 ml SR medium. 9. Under the microscope use a pipet to transfer healthy looking hESC clumps to fresh well of 6-well plate containing 2 ml of SR medium/well without feeders. Steps 8 and 9 help remove as much of the DMSO from the clumps as possible.

Derivation and Propagation of hESC Under a Therapeutic Environment

10. Transfer these clumps (5 to 10 clumps/well) into a fresh 6-well plate with HFF feeders. Incubate at 37◦ C in a humidified, 5% CO2 incubator. Take care to transfer hESC clumps in a minimum volume (∼10 to 15 hESC clumps/10 µl) to avoid carrying over DMSO.

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VITRIFICATION (FAST FREEZING) AND THAWING hESC COLONIES/CLUMPS

SUPPORT PROTOCOL 6

Vitrification is a rapid process for efficient cryopreservation of hESC. It gives a better post-thaw recovery rate of hESC compared to that in slow freezing procedure. Cryopreservation of hESC clumps at low passage number is used later for propagation and lineage specification studies. This method is adapted from ES Cell International Pty Ltd Version 2 (http://www.stemcell.edu.sg/resources/methodsProtocols.php).

Materials HEPES (Invitrogen, no. 15630-080) DMEM (Invitrogen, no. 11965-092) KOSR (Invitrogen, no. 10828-028) Sucrose Fetal bovine serum (FBS; Invitrogen, no. 16000-044) Ethylene glycol (Sigma, no. E-9129) Dimethylsulfoxide (DMSO; Sigma, D2650) hESC colonies harvested by collagenase/dispase (Support Protocol 3) Liquid nitrogen HFF feeder plates (Support Protocol 8) SR medium (see recipe) 0.22-µm syringe filter 15-ml tube Pipettor Organ culture dishes (Falcon, cat. no. 353037) for vitrification, prewarmed Open pulled straws (LEC Instruments) 5-ml cryovials with holes punched through the upper section, the bottom, and lid using a heated 18-G needle, attached to a cryostraw Liquid nitrogen tank Forceps NOTE: Wear safety glasses and gloves when working with liquid nitrogen.

Prepare vitrification solutions 1. Prepare 20.5 ml DMEM-HEPES by adding 0.5 ml of 1 M HEPES to 20 ml of DMEM. Store medium at 4◦ C. Discard any unused medium after one week. 2. Prepare 10 ml of ES-HEPES medium by adding 2 ml of KOSR to 8 ml DMEMHEPES. Prewet a 0.22-µm syringe filter with 5 ml DMEM medium. Filter the ES-HEPES solution. Store medium at 4◦ C. Discard any unused medium after one week. 3. Prepare a 1 M sucrose solution. Add 3.42 g of sucrose to 6 ml DMEM-HEPES in a 15-ml tube. Warm the solution to 37◦ C to dissolve the sucrose. If necessary, bring the solution to 8 ml with DMEM-HEPES. Add 2 ml FBS or KOSR to the solution. Filter the solution through a 0.22-µm syringe filter prewet with 5 ml DMEM. Store the solution at 4◦ C. Discard any unused solution after one week. 4. Prepare 2.5 ml of 10% vitrification solution. To 2 ml ES-HEPES add 0.25 ml ethylene glycol and 0.25 ml DMSO. Store at 4◦ C. Discard any remaining solution after each day. 5. Prepare 2.5 ml of 20% vitrification solution. To 0.75 ml ES-HEPES add 0.75 ml 1 M sucrose solution, 0.5 ml ethylene glycol, and 0.5 ml DMSO. Store at 4◦ C. Discard any remaining solution after each day.

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Freeze the clumps 6. Pick up collagenase/dispase treated hESC colonies (Support Protocol 3). Prepare pieces that are larger than those used for passaging. Larger pieces, equivalent to four pieces/hESC colony, give better post-thaw recovery

7. Using a pipettor, transfer the colony pieces to an organ culture dish containing 1 ml ES-HEPES. 8. Transfer 6 to 10 colony pieces to a second organ culture dish containing 1 ml of 10% vitrification solution for 1 min. Check that the pieces have settled to the bottom of the well. The colony pieces may “swirl” in the more viscous solution.

9. During this minute, make 10-µl drops of 20% vitrification solution on the inside of the lid of an organ culture dish, one per straw to be frozen. 10. Transfer the colony pieces to the drop of 20% vitrification solution for 25 sec. hESC are very sensitive to vitrification solutions at room temperature and extra care is required not to overexpose hESC to vitrifications solutions for time more than recommended in the step.

11. Immediately after 25-sec incubation, touch the narrow end of the vitrification straw to the side of the droplet at a 30◦ angle to the plane of the dish. The droplet should be sucked up by capillary action to make a 1-mm medium column in the straw. If this is not successful, use a pipettor on the other end of the straw to draw up pieces into it. Since working quickly is essential, it is better to leave hESC pieces in the droplet that are not picked up within the specified time behind.

12. Plunge the straw into liquid nitrogen at a 45◦ angle. 13. Transfer the straw into a labeled storage cryovial held on a cane, being careful not to push the straw into other straws already in the cryovial.

Thaw vitrified hESC clumps 14. Prepare 5 ml 0.2 M sucrose solution. To 4 ml ES-HEPES medium add 1 ml 1 M sucrose. Store at 4◦ C. Discard any remaining solution after each day. 15. Prepare 5 ml of 0.1 M sucrose solution. To 4.5 ml ES-HEPES medium, add 0.5 ml 1 M sucrose solution filter sterilized. Store at 4◦ C. Discard any remaining solution after each day. 16. Prepare a 6-well vitrification thawing plate. Add 1 ml 0.2 M sucrose solution to one well, add 1 ml 0.1 M sucrose solution to another, and 1 ml ES-HEPES medium to each of two wells. 17. Collect the cryovial containing the vitrification straws in a receptacle containing liquid nitrogen. 18. Remove a straw using forceps. Hold the straw between thumb and middle finger with the large end pointed away from your eyes to avoid liquid nitrogen that spits out. Safety glasses should be used.

Derivation and Propagation of hESC Under a Therapeutic Environment

19. Within 3 sec submerge the narrow end of the straw containing the vitrified liquid column (which contains the cell colonies) into the first well containing 0.2 M sucrose solution, at a slight angle.

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Current Protocols in Stem Cell Biology

Table 1A.4.2 Guide to the Morphology of hESC Colonies during Recovery from Freeze/Thaw

Day

Description

0

Colony pieces should appear as if they have just been cut. There should be no discernible freezing damage which may appear as: (1) bubbles attached to colony pieces, (2) floating colony pieces, (3) fragmenting colony pieces, (4) colony pieces with disintegrating patches, (5) colony pieces which are initially cohesive before disintegrating by the final thaw solution. There will be a lot of debris in the thawing plate as the colony pieces are thawed, this is normal.

1

The colony pieces tend to “disintegrate.” In this time, the healthy cells attach to the feeder layer whilst any cells damaged during freezing/thawing appear as debris in the media. It may be difficult to see attached cells so don’t panic.

2

Central “button” from colony piece becomes flatter and less distinct as cells start to grow outwards. Some colonies grow rapidly but these often become cystic.

3

Cells grow outwards but don’t show distinct colony morphology. The central button becomes less defined. More advanced colonies may appear like “targets” with an outer ring around a central denser area with an area of thin cell growth in between.

4

Small colonies should be starting to look healthy and exhibit normal morphology. Some colonies may look like small groups of cells. These will take longer to grow up and may not be the healthiest colonies.

5

Colonies should be larger but the may still appear thin.

6

Colonies starting to look healthy and thick.

7

View daily to determine when to transfer.

20. As soon as the liquid column melts place a finger on the top of the straw. As the gases in the straw expand, they should expel the liquid column from the straw. Taking your finger from the top of the straw will cause medium to move by capillary action back into the straw. If you think that the colonies may be stuck in the straw, allow medium back into the straw and then insert a 1-ml syringe fitted with a 20-µl pipet tip into the top of the straw to push the medium out.

21. After 1 min transfer the colony pieces to the next well containing 0.1 M sucrose solution. 22. After 5 min transfer the colony pieces to the next well containing ES-HEPES medium. 23. After 5 min transfer the colony pieces to the next well containing ES-HEPES medium. 24. Transfer the pieces to prepared HFF feeder 6-well plates containing 2 ml SR medium/well.

Monitor the growth of the cultures 25. Monitor the growth of the colonies. Change the medium daily. Passage the cells weekly Table 1A.4.2 is a guide to what the hESC colonies should look like after thawing. This is a rough guide only. Colonies may take longer to recover than the timeframe given here. If they do take longer, grow them to a healthy size before transfer. The colonies will look very unhealthy for the first few days but this is entirely normal. If the colony pieces exhibit no change during the first week or if they are aspirated off during media changes then it is likely that the colony pieces did not survive either the freezing or thawing process. Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 7

DERIVATION OF SERUM-FREE HUMAN FETAL FIBROBLASTS (HFF) It is estimated that ∼414 new hESC lines have been produced world-wide and only a few of these lines are characterized to some extent and available for research. The majority of these hESC lines are derived on mouse embryo fibroblasts (MEFs), but some have been derived on human-tissue-derived feeders (e.g., fetal muscle, skin, and foreskin, adult fallopian tube epithelial cells) including some under feeder-free/serumfree conditions but with undefined matrices, hence they are not suitable for clinical applications (Carpenter et al., 2004; Rosler et al., 2004; Sato et al., 2004; Beattie et al., 2005; Genbacev et al., 2005; Inzunza et al., 2005; Xu et al., 2005a; Rajan et al. 2007). This protocol describes a simple procedure on how to obtain a serum-free (KOSR contains some serum components) feeder layer for derivation and propagation of undifferentiated hESC colonies. Serum-free human fetal fibroblasts (HFF) are derived from human fetal skin after therapeutic termination of pregnancies and after obtaining informed maternal consent and institutional ethics approval. All the steps below are carried out in a standard biological safety cabinet (BSC), class II.

Materials Skin from 10- to 12-week fetuses Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen) Penicillin-streptomycin (Invitrogen) TrypLE Select (Invitrogen) SR medium (see recipe), equilibrated 35-mm petri dish Scissors 15-ml conical centrifuge tube (Fisher, cat. no. 05-539-2) Collagen type IV–coated 75-cm2 tissue culture flask (see recipe) Additional reagents and equipment for cryopreserving using a standard slow freezing procedure (Support Protocol 8) 1. Cut five to ten 2 × 3-mm2 pieces of human fetal whole skin obtained from 10- to 12-week-old fetuses after therapeutic termination of pregnancy and place them in a 35-mm petri dish. 2. Wash the pieces twice with 5 ml PBS containing 25 U/ml penicillin and 25 µg/ml streptomycin (from a stock solution purchased from Invitrogen) each time. 3. Chop the washed pieces into fine pieces, <0.1-mm, with a pair of scissors. A repeated chopping action produces almost a slurry.

4. Prepare a single-cell suspension by treating the slurry with 5 ml of TripLE Select 15 to 20 min at 37◦ C. At this stage the preparation should be a thick viscous fluid.

5. Wash the single-cell suspension in 10 ml of SR medium in a 15-ml Falcon tube by centrifuging 7 min at 500 to 600 × g, room temperature and resuspend the pellet in a fresh 10 ml SR medium. 6. Transfer the whole suspension from step 5 to a collagen type IV–coated 75-cm2 tissue culture flask containing 20 ml SR medium. Derivation and Propagation of hESC Under a Therapeutic Environment

1A.4.22 Supplement 6

The 75-cm2 tissue culture flask should be coated with 5 µg/ml human collagen IV at least 1 hr before use. Alternatively the flask could be coated with a combination of human laminin and human collagen IV (5 µg/ml each) for better adhesion of fibroblasts.

7. Incubate at 37◦ C and 5% CO2 air mixture and monitor the growth of the culture. Semi-confluent monolayer culture of HFF is visible by day 3. Current Protocols in Stem Cell Biology

8. Subculture the confluent cultures using TrypLE Select (Support Protocol 8) after 7 days and cryopreserve them by a standard slow freezing procedure in 10% DMSO (Support Protocol 5). A relatively pure population of HFF is obtained after 2 to 3 passages.

9. Validate these newly derived serum-free HFF for the ability to support the undifferentiated growth of any hESC line available before using the new HFF line for deriving new hESC lines.

PREPARING FEEDER PLATES USING HUMAN FETAL FIBROBLASTS hESC are normally grown on feeder cells for maintaining pluripotency. This protocol describes a simple procedure to prepare human fibroblast feeder layers in 6-well plates.

SUPPORT PROTOCOL 8

Materials HFF in 75-cm2 flask grown for 3 to 4 days in SR medium (see Support Protocol 7) Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen) TrypLE Select SR medium (see recipe), prewarmed Human collagen IV (Sigma) and/or human laminin 15-ml conical centrifuge tubes Benchtop centrifuge 20-µl pipettor Hemacytometer 6-well tissue culture plates (Greiner bio-one, GmbH) γ irradiator Additional reagents and equipment for counting cells (Phelan, 2006) Harvest HFF Cells 1. Wash each 75-cm2 flask containing confluent HFF monolayer with 10 ml prewarmed D-PBS twice. 2. Treat each flask with prewarmed 2 ml TrypLE Select for 2 min and vigorously shake. 3. Add 6 ml prewarmed SR medium per flask and transfer contents to a 15-ml tube. 4. Spin 7 min at ∼700 × g, room temperature. 5. Resuspend pellet in 5 ml prewarmed SR medium.

Count cells 6. Vigorously pipet cell suspension and use a sterile 20-µl pipettor to remove 10 µl of cell suspension and add directly to hemacytometer chamber. 7. Count cells in four squares of the hemacytometer (Phelan, 2006) and calculate average cell count. 8. Calculate cells per milliliter in the suspension by multiplying the average cell count by the dilution factor (if diluted) and divide by 10−4 ml. 9. Calculate the total cell number by multiplying the number of cells per ml by the total suspension volume.

Prepare 6-well feeder plates 10. Precoat the number of 6-well plates required with extracellular matrices (ECM; 1 ml/well) and keep in hood for at least 1 hr. Human collagen IV or human laminin or both together (1:1) can be used to precoat the plates. Current Protocols in Stem Cell Biology

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11. Calculate the number of cells needed to make feeder plates: the desired feeder density (1.5 × 105 cells/ml) × 2 ml/well × 6 wells/plate × number of plates. As a rule of thumb, one plate per embryo is recommended. 12. Calculate the amount of suspension to remove to give the number of cells needed by dividing the number needed for a given number of plates by the cells/ml (density of the suspension).

Irradiate HFF cells 13. Transfer the calculated volume of cells to a 15-ml conical centrifuge tube in not more than 10 ml. 14. Irradiate cells at 45 Gy for 5 to 6 min 15. Centrifuge cells 7 min at 500 to 600 × g, room temperature and resuspend in required volume of SR medium at 1.5 × 105 /ml 16. Mix the cells in the tube to distribute the cells evenly, and aliquot 2 ml/well to coated 6-well plates. SUPPORT PROTOCOL 9

FREEZING AND THAWING FROZEN HUMAN FETAL FIBROBLASTS This procedure describes how to cryopreserve HFF at low passage number. HFF are used at passage 5 as feeder layer for growing hESC. Cryopreservation of low-passage HFF is an efficient way of keeping a stock and preparing feeder layers on demand.

Materials 75-cm2 flask with HFF (Support Protocol 7) Calcium- and magnesium-free phosphate-buffered saline (D-PBS; Invitrogen), prewarmed TrypLE Select, prewarmed SR medium (see recipe), prewarmed Cryopreservation solution: 20% (v/v) DMSO in SR medium (see recipe), sterile filtered 15-ml tubes Benchtop centrifuge 0.22-µm syringe filter Cryovials Nalgene Cryofreezing containers (Fisher Scientific, Nalgene 5100-001) −80◦ C freezer Cryoboxes Liquid nitrogen tank 37◦ C water bath Freeze HFFs 1. Wash each 75-cm2 flask of low-passage HFF twice with 10 ml D-PBS. 2. Remove PBS and add 2 ml TrypLE Select. Incubate for 2 to 5 min at room temperature. 3. Add 8 ml SR medium and transfer contents to 15-ml tube. 4. Centrifuge 7 min at 700 × g (1300 rpm), room temperature. Derivation and Propagation of hESC Under a Therapeutic Environment

5. Decant the supernatant, and resuspend cells by flicking the tube. 6. Add 1.5 ml SR medium and 1.5 ml of filter sterilized cryopreservation solution slowly.

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Current Protocols in Stem Cell Biology

7. Aliquot 1 ml cells per carefully labeled cryovial immediately. Three cryovials per 75-cm2 flask are prepared.

8. Freeze at −80◦ C in Nalgene freezing container overnight. 9. Arrange cryovials in cryoboxes. 10. Store in liquid nitrogen tank.

Thaw HFFs 11. Remove a vial of frozen HFF from the liquid nitrogen tank and put it in directly into a clean water bath at 37◦ C. 12. After thawing (1 to 2 min), transfer the contents to a 15-ml tube and dilute the contents of the vial to 10 ml with SR medium. Mix thoroughly to resuspend the cells and spin 7 min at 700 × g, room temperature. 13. Decant most of the supernatant and resuspend the cells in the remaining medium in tube by vigorous shaking. Bring the volume to 5 ml with SR medium. 14. Pour the contents into 75-cm2 flask and the total volume of SR medium to 20 ml. Incubate 72 hr at 37◦ C before harvesting HFF. Change medium on an alternative day basis.

REAGENT SUPPLY AND BATCH TESTING Maintaining an adequate inventory of the reagents and their reliable suppliers are very critical for optimal functioning of the laboratory. Two types of ordering regimens are followed in our laboratory, i.e., (1) planned orders to ensure a regular supply of reagents— i.e., media, media components, and growth factors—over a period of time; and (2) periodic orders of more specialized reagents, as and when required.

SUPPORT PROTOCOL 10

Batch-to-batch variations in reagents, particularly sera (SR), extracellular matrices (ECM), and feeder cells are common and need batch testing. SR and ECM can be batch tested for cell expansion (feeders and hESC) over two passages.

Materials Human foreskin fibroblast cultures (Support Protocol 7) SR medium prepared with test and current lots of serum or serum replacement 6-well culture plates coated with test and current (control) lots of matrix Additional reagents and equipment for cell counting (Phelan, 2006) 1. Seed HFF at 1.5 × 105 /ml in SR medium with control and different test batches of serum replacement (20% SR) on plates coated with control and test lots of ECM (0.1%) in triplicate. 2. Harvest cells after 5 days. Count (Phelan, 2006) and passage the cells into fresh plates (passage 1) with the same lots of SR and/or ECM. 3. Harvest and count cells after 5 days (passage 2). Good suitable batches of SR and ECM should yield cell numbers in excess of 80% of those obtained in control cultures. Similar batch testing for sera on hESC is made by growing these cells for at least four passages. Each time assessing percentage of hESC colonies that stained positive for alkaline phosphatase (>95%), indicating the cells have retained pluripotency.

4. Similarly test new batches of HFF for the ability to support the growth of undifferentiated growth of hESC for 3 to 5 passages.

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

bFGF Add 0.1 g human serum albumin (hSA; Sigma, cat. no. A4327) to 100 ml PBS+ [with calcium and magnesium; (0.1% w/v final)]. Prewet an 0.22-µm filter with 1 ml PBS+. Filter sterilize ∼10 ml hSA solution through the filter. Aliquot 5 ml of sterile hSA solution to a sterile 14-ml centrifuge tube. Add 10 µg of bFGF (Invitrogen, cat. no. 13256-029) to 5 ml hSA solution and mix gently. Gently wash out vial containing bFGF to remove all lyophilized powder. This will give a stock concentration of 2 ng/µl.

Aliquot 0.5-ml bFGF/hSA solution to sterile microcentrifuge tubes. Label tube with concentration (2 ng/µl) and date. Store aliquots at −20◦ C (nonfrost free) or −70◦ C not more than a month.

Collagen type IV–coated culture ware Stock solution Working aseptically, prepare a 1 mg/ml stock solution of collagen type IV (Sigma, cat. no. 5533) by dissolving 5 mg in 5 ml of sterile water. Aliquot 1 ml per vial into 5 vials. Store up to several months at −20◦ C.

Working solution Prepare 5 µg/ml working solution immediately before use from the stock solution by diluting with 200 ml sterile (autoclaved) water. Coat wells and/or 75-cm2 tissue culture flasks with working solution by adding at least 1 ml per well of a 6-well plate or 3 ml for 75-cm2 flask. Tilt plate/flask in several directions to ensure that liquid covers the entire surface area. Place plates/flasks in hood for at least 1 hr or in an incubator for overnight. These coated plates/flasks can be stored for 1 week at 4◦ C. Prior to plating irradiated HFF, aspirate remaining collagen solution.

SR medium, 20% For 50 ml of the medium combine the following reagents: 37.56 ml high glucose “Knockout” DMEM (Invitrogen, cat. no. 10829-018) 0.50 ml 10 mM (100×) non-essential amino acids (NEAA; Invitrogen, cat. no 12383-014) 0.09 ml 55 mM (100×) buffered 2-mercaptoethanol 0.50 ml 200 mM (100×) L-glutamine 0.25 ml 5000 U/ml penicillin/5000 µg/mg streptomycin (Invitrogen, cat. no. 15070063) 1 ml 100× Insulin-Transferrin-Selenium (ITS; Invitrogen, cat. no. 41400-045)

Derivation and Propagation of hESC Under a Therapeutic Environment

Prewet an 0.22-µm filter (Millipore steritop) with 10 ml unsupplemented DMEM. Filter medium containing the above ingredients into a sterile 75-cm2 tissue culture flask or 50-ml Falcon tube. Add 10 ml KOSR (20%; Invitrogen, cat. no. 10828-028) to the medium after filtration and swirl gently to mix. Remove 5 ml to 25-cm2 tissue culture flask for sterility test at 37◦ C; 5% CO2 . Store medium up to 4 weeks in the dark at 4◦ C. Add 0.1 ml basic fibroblast growth factor (bFGF; see recipe) to a final concentration of 4 ng/ml to SR medium before use.

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COMMENTARY Background Information The pluripotent nature of hESCs makes them attractive as a source of various cell types that could be used for therapeutic purposes. However, eliminating all sources of contamination, animal-derived or human cell-derived, during hESC derivation and propagation is necessary before attempting the use of hESC derivatives clinically. There has been a rapid progress in this direction during the last 6 to 8 years. Following the first report of successful derivation of five hESC lines by Thomson group in 1998 (Thomson et al., 1998), it is estimated that, as of this printing, ∼414 new hESC lines have been produced world-wide, and ∼78 of these are listed in the National Institute Health (NIH) Registry (Guhr et al., 2006; Klimanskaya et al., 2006; Revazova et al., 2007; Sidhu et al., 2008). Only a few of these lines are characterized to some extent and available for research. Many of these hESC lines are not clonal, are derived under different culture conditions, and propagated on different feeder layers, the majority on mouse embryonic fibroblasts (MEFs); some lines have been derived on human tissue–derived feeders (fetal muscle, skin, and foreskin, adult fallopian tube epithelial cells) including some under feederfree/serum-free conditions which used undefined matrices, hence they are not suitable for clinical applications (Carpenter et al., 2004; Rosler et al., 2004; Sato et al., 2004; Beattie et al., 2005; Genbacev et al., 2005; Inzunza et al., 2005; Xu et al., 2005a; Rajan et al. 2007; Sidhu et al., 2008). Subtle differences in gene expression have been reported in some of these lines and in clonal lines (Richards et al., 2003; Inzunza et al., 2004; Sidhu et al., 2008). Recently some attempts have been made to derive new hESC lines in more defined conditions including serum-free or feeder-free conditions (Heins et al., 2004; Genbacev et al., 2005; Klimanskaya et al., 2005; Wang et al., 2005; Ludwig et al., 2006). However, most of these studies employed immunosurgery for dissecting inner cell masses (ICM) from embryos and they used fetal bovine serum (FBS) to grow feeder layers. Two of such hESC lines, derived recently by Ludwig et al. (2006), show chromosomal abnormalities. Similarly Ellerstrom and coworkers (2006) recently tried defining an in vitro culture system for the derivation of a new hESC line under xenofree conditions using human serum that also caused differentiation of hESC in long-term cultures. Some biotech companies are now

offering defined culturing kits for propagation of hESC under feeder-free, xeno-free, and serum-free environments (Invitrogen, Millipore, and BD); these are yet to be validated in different laboratories. While most common contaminations in tissue culture laboratory environment (e.g., mycoplasma, bacteria, yeast, and fungi) can be minimized, the use of sera and feeders imposes serious virological risks (both murine-type, i.e., LCMV, reovirus-3, and human-types, i.e., rabies SARS, HTLV-3, 4). There are also viral risks associated with the use of serum (e.g., HIV 1 and 2, hepatitis B, C). Eliminating such risks becomes mandatory if hESC derivatives are to be used for therapeutic purposes. Accredited stem cell banks can offer valuable support in adopting standardized operating procedures and safety measures for validation, quality, and safety of new hESC produced (Stacey et al., 2006). For a summary of the hESC lines derived so far, see Table 1A.4.1.

Critical Parameters and Troubleshooting GMP compliance Maintaining GMP compliance according to the regulatory authority’s guidelines is very critical for accreditation of the final product. Four main parameters for keeping GMP compliance—i.e., standard operating procedures, reagent supply and batch testing, maintaining stocks, cell banking and distribution, and work/time flow for derivation of hESC lines—need to be followed strictly in order to avoid any trouble in GMP compliance. Keeping a weekly log of these activities is very helpful in troubleshooting. An audit of GMP facility every 6 months by professional agencies is also helpful before the main audit by the regulatory authority. Thawing and culture of embryos This is normally carried out by an embryologist. Record keeping of all the donated embryos is critical from ethics points of view and is also relevant for back reference if a new hESC is established. Early stage embryos are cultured in the specified medium for development to the blastocyst stage. The later stage embryos normally hatch after further incubation for 24 hr. If these blastocysts don’t hatch in 24 hr, a gentle zona breaching by using a dissecting pipet or laser is recommended. The hatched blastocysts are allowed to attach

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to the feeder layer before attempting ICM isolation. Dissection of inner cell mass and co-culture on feeder Dissection of ICM is recommended after attachment of the blastocyst to the feeder layer within 3 to 4 days of culture. If blastocysts are not fully spread out and the ICM is not clearly visible, another day of culture may be carried out. Dissection of ICM by laser is a quick process only if ICM is clearly visible. If ICM is not clearly visible, whole embryo culture is recommended. Establishing a new hESC line This is a sequential process involving many steps. The critical parameters and troubleshooting for each of these steps are described below. Passaging hESC with collagenase/dispase Table 1A.4.3 provides troubleshooting information for passaging hESC colonies with collagenase/dispase (Support Protocol 3).

Isolation of HFFs Aseptic conditions must be followed throughout isolation of HFFs, and a BSC class II hood is recommended for processing the tissue. Batch testing for ECM (collagen IV/laminin) and KOSR is recommended (see Support Protocol 10). Digestion of the tissue pieces with TrypLE Select (Support Protocol 4) for 15 to 20 min should yield viscous slurry from tissue but if it does not, increase the time of incubation up to 30 min. Table 1A.4.4 provides troubleshooting information for isolation of HFFs (Support Protocol 7). Preparing HFF feeder plates Table 1A.4.5 provides troubleshooting information for preparation of HFF feeder plates (Support Protocol 8). Freezing and thawing HFF DMSO is toxic to cells at room temperature and care should be taken to chill the

Table 1A.4.3 Troubleshooting Guide to Passaging hESC Colonies with Collagenase/Dispase

Symptoms

Possible causes

Comments

hESC single cells/clumps are nonviable

1. hESC are overexposed to collagenase

1. Optimize collagenase/TrypLE Select treatment time or use as recommended

2. hESC single cells don’t survive very well in culture medium alone

2. Use HFF-conditioned medium (24 hr) for hESC single cells as supplement to SR medium

Table 1A.4.4 Troubleshooting Guide to Isolation of Human Fetal Fibroblasts

Symptoms

Possible causes

Comments

HFF do not attach to 75-cm2 flask or form islands

1. Flasks surface is not uniformly coated with ECM

1. Make sure enough of ECM in solution (3-4 ml) covers the whole surface of the flask at least for 1 hr at room temperature

2. HFF are treated with TrypLE Select longer than recommended

2. Reduce the time of digestion with TrypLE Select

3. bFGF is not added to SR medium

3. Add bFGF to SR medium

1. bFGF is not added to SR medium

1. Add bFGF to SR medium

Slow growing HFF Derivation and Propagation of hESC Under a Therapeutic Environment

2. Human skin tissue was 2. Use fresh human fetal skin tissue that is from fetuses not transported transported quickly within 1-2 hr from quickly to the laboratory and clinics hence stale

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Table 1A.4.5 Troubleshooting Guide to Preparing HFF Feeder Plates

Symptoms

Possible Causes

Comments

HFF are not uniformly distributed and form islands

1. The 6-well tissue culture 1. Make sure enough of ECM in solution plate surface is not uniformly (1 ml/well) covers the whole surface of the coated with ECM well at least for 1 hr at room temperature 2. HFF numbers are not as recommended

2. Seed HFF at 1.5 × 105 /ml

3. HFF are overexposed to γ-irradiation

3. HFF are overexposed to γ-irradiation (the optimal exposure recommended is 45 Gy/7 min) 4. Alternately treat HFF with mitomycin C (10 µg/ml) for 2 hr

cryovials on ice during the freezing process for HFF in a BSC Class II hood (Support Protocol 9). HFF tend to die if stored for an extended period (>2 to 3 days) at −80◦ C. The frozen vials from Nalgene container should be transferred to liquid nitrogen after the overnight incubation.

Anticipated Results The protocols described for producing hESC lines in this unit ensure maintenance of quality control that is essential for GMP compliance. The success of obtaining hESC lines depends on many factors, primary being the quality of donated embryos, culturing conditions, and handling procedures. The success rate for isolating a new stem cell line varies from 5% to 20%.

Time Considerations From hatching of blastocyst to its attachment onto feeder layer and the appearance of the first hESC colony takes ∼10 days. The first hESC colony is physically dissected into 6 to 8 pieces and transferred to fresh feeder plate. Once 10 to 15 good looking hESC colonies are obtained, these can be passaged (mechanical passaging, UNIT 1C.1) into a new 6-well plate. It takes ∼1 week to go from one 6-well plate to six new 6-well plate cultures. Freezing of an aliquot at this stage is strongly recommended. hESC colonies from each 6-well plate can be transferred now by using TrypLE Select (see Support Protocol 4) to three 75-cm2 flasks. Within 3 weeks eighteen new 75-cm2 flasks containing hESC can be produced. Regular freezing and characterization can be carried out at this stage. Regular weekly passage is very essential to maintain pluripotency in hESC.

Acknowledgements Dr. Kuet Li served as a consultant for the overall strategy of the GMP facility and the requirements for obtaining an Australian license for preparing hESC under GMP. Dr. Sidhu produced the protocols on hESCs while at the Diabetes Transplant Unit, Prince of Wales Hospital, University of New South Wales. These protocols are reproduced with permission of the director.

Literature Cited Amit, M., Shariki, C., Margulets, V., and ItskovitzEldor, J. 2004. Feeder layer and serum-free culture of human embryonic stem cells. Biol. Reprod. 70:837-845. Beattie, G.M., Lopez, A.D., Bucay, N., Hinton, A., Firpo, M.T., King, C.C., and Hayek, A. 2005. Activin A maintains pluripotency of human embryonic stem cells in the absence of feeder layers. Stem Cells 23:489-495. Brimble, S.N., Zeng, X., Weiler, D.A., Luo, Y, Liu, Y, Lyons, I.G., Freed, W.J., Robins, A.J., Rao, M.S., and Schulz, T.C. 2004. Karyotypic stability, genotyping, differentiation, feeder-free maintenance, and gene expression sampling in three human embryonic stem cell lines derived prior to August 9, 2001. Stem Cells Dev. 13:585597. Carpenter, M.K., Rosler, E.S., Fisk, G.J., Brandenberger, R., Ares, X., Miura, T., Lucero, M., and Rao, M.S. 2004. Properties of four human embryonic stem cell lines maintained in a feederfree culture system. Dev. Dynamics 229:243253. Cheng, L., Hammond, H., Ye, Z., Zhan, X., and Dravid, G. 2003. Human adult marrow cells support prolonged expansion of human embryonic stem cells in culture. Stem Cells 21:131-142. Draper, J.S., Smith, K., Gokhale, P., Moore, H.D., Maltby, E., Johnson, J., Meisner, L., Zwaka, T.P., Thomson, J.A., and Andrews, P.W. 2004. Recurrent gain of chromosomes 17q and 12

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in cultured human embryonic stem cells. Nat. Biotechnol. 22:53-54. Ellerstrom, C., Strehl, R., Moya, K., Andersson, K., Bergh, C., Lundin, K., Hyllner, J., and Semb, H. 2006. Derivation of a xeno-free human embryonic stem cell line. Stem Cells 24:2170-2176. Genbacev, O., Krtolica, A., Zdravkovic, T., Brunette, E., Powell, S., Nath, A., Caceres, E., McMaster, M., McDonagh, S., Li, Y., Mandalam, R., Lebkowski, J., and Fisher, S.J. 2005. Serum-free derivation of human embryonic stem cell lines on human placental fibroblast feeders. Fertil. Steril. 83:1517-1529. Guhr, A., Kurtz, A., Friedgen, K., and Loser, P. 2006. Current state of human embryonic stem cell research: An overview of cell lines and their use in experimental world. Stem Cells 24:21872191. Heins, N., Englund, M.C.O., Sjoblom, C., Dahi, U., Tonning, A., Bergh, C., Lindahl, A., Hanson, C., and Semb, H. 2004. Derivation, characterization, and differentiation of human embryonic stem cells. Stem Cells 22:367-376. Hovatta, O., Mikkola, M., Gertow, K., Stromberg, A.M., Inzunza, J., Hreinsson, J., Rozell, B., Blennow, E., Andang, M., and Ahrlund-Richter, L. A. 2003. Culture system using human foreskin fibroblasts as feeder cells allows production of human embryonic stem cells. Hum. Reprod. 18:1404-1409. Inzunza, J., Gertow, K., Stromberg, M.A., Matilainen, E., Blennow, E., Skottman, H., Wolbank, S., Ahrlund-Richter, L., and Hovatta, O. 2005. Derivation of human embryonic stem cell lines in serum replacement medium using postnatal human fibroblasts as feeder cells. Stem Cells 23:544-549. Inzunza, J., Sahlen, S., Holmberg, K., Stromberg, A.M., Teerijoki, H., Blennow, E., Hovatta, O., and Malmgren, H. 2004. Comparative genomic hybridization and karyotyping of human embryonic stem cells reveals the occurrence of an isodicentric X chromosome after long-term cultivation. Mol. Hum. Reprod. 10:461-466.

Derivation and Propagation of hESC Under a Therapeutic Environment

man embryonic stem cells in defined serumfree medium devoid of animal-derived products. Biotechnol. Bioeng. 91:688-698. Ludwig, T.E., Levenstein, M.E., Jones, J.M., Berggren, W.T., Mitchen, E.R., Frane, J.L., Crandall, L.J., Daigh, C.A., Conard, K.R., Piekarczyk, M.S., Llanas, R.A., and Thomson, J.A. 2006. Derivation of human embryonic stem cells in defined conditions. Nat. Biotechnol. 24:185-187. Mallon, B.S., Park, K.Y., Chen, K.G., Hamilton, R.S., and McKay, R.D. 2006. Toward xeno-free culture of human embryonic stem cells. Int. J. Biochem. Cell Biol. 38:1063-1075. Miyamoto, K., Hayashi, K., Suzuki, T., Ichihara, S., Yamada, T., Kano, Y., Yamabe, T., and Ito, Y. 2004. Human placenta feeder layers support undifferentiated growth of primate embryonic stem cells. Stem Cells 22:433-40. Park, S.P., Lee, Y.J., Lee, K.S., Shin, A. H., Cho, H.Y., Chung, K.S., Kim, E.Y., and Lim, J.H. 2004. Establishment of human embryonic stem cell lines from frozen-thawed blastocysts using STO cell feeder layers. Hum. Reprod. 19:67684. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Rajan, P., Smotrich, D., Ross, R., Larent, L., and Loring, J.F. 2007. Derivation of embryonic stem cells from human blastocysts. In Human Stem Cell Manual a Laboratory Guide (J.F. Loring, R.L. Wesselschmidt, and P.H. Schwartz eds.). Academic Press, N.Y. Revazova, E.S., Turovets, N.A., Kochetkova, O.D., Kindarova, L.B., Kuzmichev, L.N., Janus, J.D., and Pryzhkova, M.V. 2007. Patient-specific stem cell lines derived from human parthenogenetic blastocysts. Cloning and Stem Cells 9:1-18. Richards, M., Fong, C. Y., Chan, W. K., Wong, P. C., and Bongso, A. 2002. Human feeders support prolonged growth of human inner cell masses and embryonic stem cells. Nat. Biotechnol. 20:933-936.

James, D., Levine, A.J., Besser, D., and HemmatiBrivanlou, A. 2005. TGFβ/activin/nodal signaling is necessary for the maintenance of pluripotency in human embryonic stem cells. Development 132:1273-1282.

Richards, M., Tan, S., Fong, C.Y., Biswas, A., Chan, W.K., and Bongso, A. 2003 Comparative evaluation of various human feeders for prolonged undifferentiated growth of human embryonic stem cells. Stem Cells 21:546-556.

Kim, H.S., Oh, S.K., Park, Y.B., Ahn, H.J., Sung, K.C., Kang, M.J., Lee, L.A., Suh, C.S., Kim, S.H., Kim, D.W., and Moon, S.Y. 2005. Methods for derivation of human embryonic stem cells. Stem Cells 23:1228-1233.

Rosler, E.S., Fisk, G.J., Ares, X., Irring, J., Miura, T., Rao, M.S., and Carpenter, M.K. 2004. Longterm culture of human embryonic stem cells in feeder-free conditions. Dev. Dynamics 229:259274.

Klimanskaya, I., Chung, Y., Meisner, L., Johnson, J., West, M.D., and Lanza, R. 2005. Human embryonic stem cells derived without feeder cells. Lancet 365:1636-1641.

Sato, N., Meijer, L., Skaltsounis, L., Greengard, P., and Brivanlou, A.H. 2004. Maintenance of pluripotency in human and mouse embryonic stem cells through activation of WNT signaling by a pharmacological GSK-3-specific inhibitor. Nat. Med. 10:55-63.

Klimanskaya, I., Chung, Y., Becker, S., Lu, S.J., and Lanza, R. 2006. Human embryonic stem cell lines derived from single blastomeres. Nature (letter) 444:481-485. Li, Y., Powell, S., Brunette, E., Lebkowski, J., and Mandalam, R. 2005. Expansion of hu-

Sidhu, K.S. and Tuch, B. E. 2006. Derivation of three clones from human embryonic stem cell lines by FACS sorting and their characterization. Stem Cells Devel. 15:61-69.

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Sidhu, K.S., Ryan, J.P., and Tuch, B.E. 2008. Derivation of a new hESC line, endeavour-1 and its clonal propagation. Stem Cells Devel. 17:4152.

Wang, Q., Fang, Z.F., Jin, F., Lu, Y., Gai, H., and Sheng, H.Z. 2005. Derivation and growing human embryonic stem cells on feeders derived from themselves. Stem Cells 23:1221-1227.

Stacey, G.N., Cobo, F., Nieto, A., Talavera, P., Healy, L., and Concha, A. 2006. The development of ‘feeder’ cells for the preparation of clinical grade hES cell lines: Challenges and solutions. J. Biotechnol. 125:583-588.

Xu, C., Inokuma, M.S., Denham, J., Golds, K., Kundu, P., Gold, J.D., and Carpenter, M.K.. 2001. Feeder-free growth of undifferentiated human embryonic stem cells. Nat. Biotechnol. 19:971-974.

Stojkovic, P., Lako, M., Przyborski, S., Stewart, R., Armstrong, L., Evans, J., Zhang, X., and Stojkovic, M. 2005a. Human-serum matrix supports undifferentiated growth of human embryonic stem cells. Stem Cells 23:895-902.

Xu, C., Jiang, J., Sottile, V., McWhir, J., Lebkowski, J., and Carpenter, M. K. 2004. Immortalized fibroblast-like cells derived from human embryonic stem cells support undifferentiated cell growth. Stem Cells 22:972-980.

Stojkovic, P., Lako, M., Stewart, R., Pryzborski, S., Armstrong, L., Evans, J., Murdoch, A., Strachan, T., and Stojkovic, M. 2005b. An autogeneic feeder cell system that efficiently supports growth of undifferentiated human embryonic stem cells. Stem Cells 23:306-314.

Xu, C., Rosler, E., Jiang, J., Lebkowski, J. S., Gold, J. D., O’Sullivan, C., Delevan-Boorsma, K., Mok, M., Bronstein, A., and Carpenter, M.K. 2005a. Basic fibroblast growth factor supports undifferentiated human embryonic stem cell growth without conditioned medium. Stem Cells 23:315-323.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshal, V.S., and Jones, J.M. 1998. Embryonic stem cell line from human blastocysts Science 282:11451147. Vallier, L., Alexander, M., and Pederson, R. A. 2005. Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell Sci. 118:4495-4509.

Xu, R. H., Peck, R. M., Li, D. S., Feng, X., Ludwig, T., and Thomson, J. A. 2005b. Basic FGF and suppression of BMP signaling sustain undifferentiated proliferation of human ES cells. Nat. Methods 2:185-190.

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Proteomic Analysis of Pluripotent Stem Cells

UNIT 1B.1

Sean C. Bendall,1 Aaron T. Booy,1 and Gilles Lajoie1 1

University of Western Ontario, London, Ontario, Canada

ABSTRACT Mass spectrometry (MS)–based proteomics has become one of the most powerful tools for identifying expressed proteins, providing quick insights into molecular and cellular biology. Traditionally, proteins isolated by either one- or two-dimensional gel electrophoresis are digested with a site specific protease. The resulting peptides are subject to one of two forms of analysis: (1) matrix-assisted laser desorption/ionization time-offlight (MALDI-TOF) MS, where a “mass fingerprint” of all the peptides in a sample is generated, or (2) electrospray ionization tandem MS (ESI-MS/MS), where a mass fragmentation spectra is generated for each peptide in a sample. The resulting mass information is then compared to that of a theoretical database created with available genomic sequence information. This unit provides protocols for this type of assessment C 2007 in embryonic stem cells (ESCs). Curr. Protoc. Stem Cell Biol. 2:1B.1.1-1B.1.33.
by John Wiley & Sons, Inc. Keywords: human embryonic stem cells r proteomics r mass spectrometry r gel electrophoresis r protein digestion r protein sequencing

INTRODUCTION Mass spectrometry (MS)–based proteomics has become one of the most powerful tools for identifying expressed proteins, providing quick insights into molecular and cellular biology (Aebersold and Mann, 2003; Steen and Mann, 2004; Domon and Aebersold, 2006). Traditionally, proteins isolated by either one- or two-dimensional gel electrophoresis are digested with a site specific protease. The resulting peptides are subject to one of two forms of analysis: (1) matrix assisted-laser desorption/ionization time-of-flight (MALDITOF) MS, where a “mass fingerprint” of all the peptides in a sample is generated, or (2) electrospray ionization tandem MS (ESI-MS/MS), where a mass fragmentation spectrum is generated for each peptide in a sample. The resulting mass information is then compared to that of a theoretical database created with available genomic sequence information. This proteomic schema has rapidly evolved, and now the ability to identify proteins based on accurate mass measurements has impacted many areas of cell biology, including: 1. Proteome characterization. This characterizes all proteins present in different biological tissues, fluids (Adachi et al., 2006), or subcellular compartments (Andersen and Mann, 2006; Foster et al., 2006). The breadth of these endeavors has recently expanded to encompass the proteome of entire organisms (Kislinger et al., 2006). 2. Functional proteomics. An alternative to the yeast two-hybrid system (e.g., see Golemis et al., 1998), this approach has been used to formulate complex protein interaction networks (Gavin et al., 2002; Ho et al., 2002; Stelzl et al., 2005). It has also been employed in the elucidation of interaction profiles between proteins and other macromolecules. Characterization of Embryonic Stem Cells Current Protocols in Stem Cell Biology 1B.1.1-1B.1.33 Published online July 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b01s2 C 2007 John Wiley & Sons, Inc. Copyright


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3. Quantitative proteomics. MS is not strictly quantitative; however, by incorporation of isotopic labels through either metabolic (Ong et al., 2002; Mann, 2006) or chemical (Ong and Mann, 2005) means, the relative abundance of proteins can be determined. Another quantitative approach is to incorporate a sample with a known amount of isotopically coded standard (Kirkpatrick et al., 2005b) to obtain more accurate values. 4. Identification of post-translational modifications (PTM). A number of MS-based strategies have been developed to identify protein PTMs (Mann and Jensen, 2003). Primarily, these strategies focused on identification of phosphorylation sites in cell signaling studies (Ptacek and Snyder, 2006; Schmelzle and White, 2006), but they have also been used successfully to characterize sites of methylation/acetylation (Beck et al., 2006), ubiquitination (Kirkpatrick et al., 2005a), and complex patterns of glycosylation (Raman et al., 2005). 5. Combinatorial proteomic approaches. The most noteworthy examples of this approach involve the combination of both quantitative and functional proteomic applications to study the dynamics of multiple protein interactions in unison (Ranish et al., 2003; Andersen et al., 2005). Others have invoked the combination of quantitative proteomics and phosphorylation analysis to both identify temporal changes in cell signaling (Blagoev et al., 2004; Blagoev and Mann, 2006) and compare/contrast signaling cascades in stem cell populations (Kratchmarova et al., 2005). To date, successful large-scale application of proteomic technologies in the embryonic stem cell system has been limited to cell lines derived from the mouse. Recent work with embryonic stem cells (ESCs) from the mouse involved an investigation of the Nanog protein-interaction regulatory network (Wang et al., 2006), which illustrated the potential these methodologies include for deciphering the uniquely complex biology of ESCs. There is essentially no barrier between many of these aforementioned technologies and application to hESCs. Many were developed using standard tissue culture cell lines, which are almost biochemically identical to ESCs. Consequently, few methodological changes are necessary to adapt current proteomic applications to hESCs. With this in mind, there are a few aspects of hESC biology that do need to be taken into consideration. Some experiments, including those investigating phosphorylation, can require up to 109 cells to be completed successfully. Unlike standard tissue culture and as described in this chapter, hESCs grow slowly and are susceptible to differentiation in culture. Consequently, it may be difficult to obtain the number of cells necessary for an experiment; this is compounded with the fact that hESCs often require sorting based on phenotypic markers in order to obtain enriched cell populations. The analytical instrumentation used in proteomics analysis is expensive and highly specialized, varying in type as much as in potential uses. Consequently, creating a generic protocol for proteomic analysis is beyond the scope of this unit. As such, the authors recommend that the researcher choose a particular proteomic application on which to model specific hESC-based experiments and pay particular attention to the MS-based approach that was employed. Prior to embarking on any investigation, consult with an MS collaborator, core facility, or service center about time, expense, and their ability to address the study’s technical needs.

Proteomic Analysis of Pluripotent Stem Cells

The protocols in this unit are designed to provide a framework for the preparation of hESCs intended for any of the aforementioned proteomic investigations. They represent a subset of protocols which are nearly universal in proteomic applications and are performed independent of the analytical instrumentation. The unit describes a variety of basic extraction protocols for obtaining hESCs proteins under denaturing conditions (Basic Protocol 1) as well as the preparation of samples for two-dimensional (2-D) gel

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electrophoresis (Alternate Protocol 1) and affinity purification applications (Alternate Protocol 2). In addition, the unit describes the basic subcellular fractioning of hESCs (Alternate Protocol 3) and collection of proteins excreted in hESC culture (Basic Protocol 2). Also detailed is quantification of protein in these extracts (Basic Protocol 3 and Alternate Protocol 4), separation by gel electrophoresis (Basic Protocol 4), and visualization by colloidal Coomassie staining (Support Protocol 2). Once separated, proteins can be digested with trypsin from a gel (Basic Protocol 5) or in solution (Alternate Protocol 5). The concentration of protein and peptide solutions and the subsequent removal of interfering substances is described in Support Protocol 1 and Basic Protocol 6, respectively. Basic Protocol 7 describes how the resulting tryptic peptides can be prefractionated by cation exchange chromatography prior to MS analysis. NOTE: Human keratin (from the dust in the air and the experimenters) is a common contaminant that arises in all aspects of proteomic sample handling. To minimize the occurrence of human keratin in samples consider the following precautions: 1. Use only freshly prepared reagents and pass all solutions through 0.22-µm filters. 2. Wear gloves and a lab coat. 3. Perform all work in a biosafety cabinet or laminar flow hood. 4. Keep all gels and samples covered; minimize handling. 5. Use disposable plastic containers and micropipet tips that are packaged by the manufacturer. Items that are washed, autoclaved, and reused can be sterile, but they are still usually contaminated with foreign protein. NOTE: Regardless of the type of proteomic experimentation, the importance of washing hESCs multiple times with PBS prior to protein extraction cannot be stressed enough. hESC culture medium contains extremely high concentrations of serum proteins. Contamination with these serum proteins can mask the signal from endogenous proteins of interest if not removed.

EXTRACTION OF PROTEIN FROM hESCs UNDER DENATURING CONDITIONS

BASIC PROTOCOL 1

The following is not meant to be a definitive protocol, but rather a guideline for successful hESC protein extraction. The resulting extracts are compatible with most popular proteomics methods. Depending on the line of experimentation and the nature of the proteomic query, alternate protein extraction and buffer composition may be necessary. Where applicable, the following sections will contain a reagents compatibility table. Those values can be used to modify procedures leading up to proteomic analysis. The authors have found that hESCs grown on matrigel in the absence of feeder cells yielded ∼50 µg of protein per 106 cells lysed. The number of cells required can be estimated, depending on the demands of the experiment. However a small-scale preparation ahead of time to properly judge efficiency and compatibility of the hESC culture system with the analysis procedures for the extracts is always recommended.

Materials Adherent hESC growing on plates or fresh or frozen (up to 6 months at −30◦ C) hESC cell pellets washed with PBS (Invitrogen; see Section C in this book for cell culture and preparation information), 106 or more cells per extraction Phosphate-buffered saline, pH 7.4 (PBS; Invitrogen), 4◦ C Denaturing cell lysis buffer (see recipe)

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1× Laemmli buffer (see recipe) Phosphatase inhibitor cocktail (see recipe), optional Protease inhibitor cocktail (see recipe), optional 20× nuclease cocktail (see recipe), optional 15-ml centrifuge tube 1-ml syringe with 22-G needle, optional Refrigerated centrifuge Lyse cells growing in plates 1a. Wash adherent hESCs three times with 300 µl/cm2 plate area, using cold (4◦ C) PBS For example, use 3 ml per wash for a 6-well plate (9.8 cm2 /well).

2a. Add enough denaturing cell lysis buffer to the plate containing the cells to just cover the surface. Let stand 5 to 10 min on ice with occasional mixing. Use a cell scraper to assist in removal of the cells. To minimize volume, if harvesting more than one well for a given sample, harvest one well at a time using the same aliquot of lysis buffer to harvest the subsequent wells. For analysis of protein phosphorylation or concerns regarding proteolytic degradation, add phosphatase and protease inhibitors prior to cell lysis.

Lyse fresh or frozen cell pellets 1b. For immediate SDS-PAGE analysis: Thaw frozen hESC pellets on ice. Lyse fresh or frozen hESC pellets directly in 25 µl/106 cells 1× Laemmli buffer, immediately heat to 65◦ C (to prevent sample degradation), and proceed with SDS-PAGE (see Basic Protocol 4). 106 cells will yield ∼2 µg protein.

2b. For other analyses: Resuspend the cell pellet in denaturing cell lysis buffer (typically 25 µl of buffer per 106 cells). Let stand on ice 10 to 20 min with occasional vortex mixing. For later proteomic analysis it is desirable here to keep volumes as low as possible to keep protein concentrations high. Buffers containing urea should be prepared just prior to use, or frozen immediately at −80◦ C after preparation to avoid protein carbamylation, which can interfere with proteomic analysis (McCarthy et al., 2003).

Extract proteins 3. Repeatedly vortex and/or mix the suspension during step 2a or 2b with the 1-ml syringe and needle until the solution begins to clear. Rest suspension on ice when not mixing. 4. Optional, to reduce foaming: Centrifuge 1 min at 5000 × g, 4◦ C, and continue to mix. 5. Optional, to reduce viscosity: Add 20× nuclease cocktail to a final concentration of 1× and continue to mix. At this point the solution may be very viscous due to the presence of unsheared genomic DNA.

6. Centrifuge the lysed cells 10 min at 10,000 × g, 4◦ C, and transfer the supernatant to another clean tube. Proteomic Analysis of Pluripotent Stem Cells

There may be a small pellet (<15% total protein) remaining in the original tube consisting of membrane and other hydrophobic proteins.

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7. Discard the pellet or, alternatively, add an equal volume of denaturing cell lysis buffer containing 2% SDS to the pellet, vortex, and then centrifuge 10 min at 10,000 × g, 4◦ C. This fraction can be combined with the “soluble” supernatant from step 6 or kept as a separate fraction, typically termed “membrane.” For further insight into options on membrane protein analysis, see (Wu et al., 2003; Wu and Yates, 2003).

8. Store the preparations on ice for use the same day or store up to 1 year at −30◦ C (or lower). To minimize freeze-thaw cycles, ∼20 µl of the preparations can be kept in separate tubes for determination of protein concentration.

EXTRACTION OF PROTEIN FOR TWO-DIMENSIONAL GEL ELECTROPHORESIS

ALTERNATE PROTOCOL 1

This protocol has been designed for extraction of protein from hESC cell pellets while leaving the extracts compatible with two-dimensional (2-D) gel electrophoresis. Protein extracted by this method is also compatible with other isoelectric focusing applications, free-flow electrophoresis, and SDS-PAGE analysis.

Additional Materials (also see Basic Protocol 1) Two-dimensional (2-D) gel extraction buffer (see recipe) Follow Basic Protocol 1 with exceptions to the following steps: 1a or 1b. Replace the denaturing cell lysis buffer with 2-D gel extraction buffer. Use of EDTA, phosphatase inhibitors, or nuclease cocktail will contribute to the ionic strength of the extract and can interfere with isoelectric focusing. Keep samples on ice and use mechanical shearing to remove DNA/RNA to avoid using inhibitors and nucleases.

7. Do not extract protein from the membrane pellet. Discard the pellet. These membrane proteins generally run very poorly in 2-D gels and should be disregarded for this application.

EXTRACTION OF PROTEIN FROM WHOLE CELLS FOR AFFINITY PURIFICATION

ALTERNATE PROTOCOL 2

This protocol has been optimized from total cell protein extraction under conditions that will still support the majority of affinity applications (i.e., protein-macromolecule interactions studies). Cell lysates created using this procedure have been used successfully in analyzing protein complexes via tandem affinity purification (TAP) tag- or antibodybased strategies. It has also proven successful in the purification of phosphorylated proteins for MS analysis using phospho-specific antibodies. While this lysis buffer (affinity extraction buffer) does contain detergent, if samples are washed with a buffer such as PBS during purification, the samples can be considered detergent free.

Additional Materials (also see Basic Protocol 1) Affinity extraction buffer (see recipe) Follow Basic Protocol 1 with exceptions to the following steps: 1a or 1b. Replace the denaturing cell lysis buffer with up to 1 ml affinity extraction buffer per 106 cells.

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Because protein extracted here is destined for affinity purification (effectively a concentration step) volume is not as much of a concern. Consequently, to increase lysis efficiency use up to 1 ml of affinity extraction buffer per 106 cells.

7. Do not extract protein from the membrane pellet. Discard the pellet. Store samples on ice or at −30◦ C (or lower) for longer periods. The use of a stronger reagent is not an option for affinity purification and consequently this fraction should not be included in further analysis. For purifications of protein complexes it is recommended that samples be prepared fresh and not frozen. ALTERNATE PROTOCOL 3

HYPOTONIC PROTEIN EXTRACTION WITH CELLULAR FRACTIONATION This procedure is useful for prefractionation of protein samples at the cellular level. It separates hESC samples into “nuclei,” “cytoplasmic,” and “membrane” fractions. For this method, hESCs need to be intact and prepared fresh. All proteins are isolated under native conditions and are still suitable for interaction studies or affinity purification. While this procedure provides a basic level of subcellular fractionation, it is inefficient and requires a relatively large number of cells. Alternatively, there are commercially available kits (e.g., Nuclear Extract kit; Active Motif) which can also generate similar fractionations.

Materials 107 to 108 fresh hESC growing on tissue culture plates or as a pellet (see Section C in this book for cell culture and preparation information) Phosphate-buffered saline, pH 7.4 (PBS; Invitrogen), prewarmed to 37◦ C Hypotonic cell lysis buffer (see recipe) Phosphatase inhibitor cocktail (see recipe) Protease inhibitor cocktail (see recipe) 40-ml Dounce homogenizer (glass grinder) 15-ml plastic, conical centrifuge tubes, precoded Temperature-controlled centrifuge Additional reagents and equipment for concentrating proteins (see Basic Protocol 2 or Support Protocol 1) and extracting proteins from nuclei and membrane fractions (see Basic Protocol 1 or Alternate Protocol 1 or 2) Lyse cells growing in plates 1a. Wash adherent hESCs three times with 300 µl/cm2 plate area, using cold (4◦ C) PBS. For example, use 3 ml per wash for a 6-well plate (9.8 cm2 /well).

2a. Add enough hypotonic cell lysis buffer to the plate containing the cells to just cover the surface. Equilibrate 20 min on ice, allowing cells to swell. Use a cell scraper to assist in removal of the cells and collect them in a precooled 15-ml conical centrifuge tube. To minimize volume, if harvesting more than one well for a given sample, harvest one well at a time using the same aliquot of lysis buffer to harvest the subsequent wells. For analysis of protein phosphorylation, or concerns regarding proteolytic degradation, add phosphatase and protease inhibitors prior to cell lysis. Proteomic Analysis of Pluripotent Stem Cells

Lyse cell pellets 1b. Wash a fresh hESC pellet with PBS (see Section C in this book).

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2b. On ice, resuspend a fresh hESC pellet in hypotonic lysis buffer by gently mixing. Use ∼10 ml for 108 cells. Equilibrate for 20 min on ice, allowing cells to swell. 3. Treat the cells with ∼20 strokes from a 40-ml Dounce homogenizer. 4. Check a sample under a microscope to verify that outer membranes have been removed form the hESCs. If a majority of cells are still intact repeat step 3 and check again.

Fractionate the sample 5. Transfer to a 15-ml centrifuge tube and centrifuge 20 min at 900 × g, 4◦ C. 6. Transfer the supernatant to a clean 15-ml tube and store on ice. 7. Resuspend the pellet in the hypotonic cell lysis buffer (∼10 ml) by gently mixing and centrifuge 20 min at 900 × g, 4◦ C. 8. Discard the supernatant. Hold the pellet (the “nuclei" fraction) on ice. 9. Centrifuge the original supernatant (from step 6) ∼3 hr at 100,000 × g, 4◦ C. The resulting supernatant from this centrifugation is the “cytosolic" fraction and the pellet is the “membrane fraction”.

10. If necessary (i.e., sample contains <1 mg/ml of protein), concentrate the cytosolic fraction, using an MWCO filter as described in Basic Protocol 2 or protein precipitation as described in Support Protocol 1. 11. Extract the proteins from both the nuclei and membrane fractions as described in Basic Protocol 1, step 7, or Alternate Protocol 1 or 2, depending on the desired application.

COLLECTION OF EXTRACELLULAR PROTEINS FROM hESC CULTURE This protocol is for the collection of proteins secreted by hESCs in culture. It has been employed for hESCs grown independent of feeder cells. In the authors’ experience, the following procedure will generate ∼1 µg of protein per ml of medium conditioned.

BASIC PROTOCOL 2

Materials Day 4 or 5 hESC on matrigel (see Xu et al., 2001; Hoffman and Carpenter, 2005; Wang et al., 2005), ∼60% to 80% confluent Sterile tissue-culture phosphate-buffered saline, pH 7.4 (PBS; Invitrogen), prewarmed to 37◦ C Protein-free hESC medium (see recipe), prewarmed to 37◦ C Denaturing cell lysis buffer (see recipe) 0.22-µm sterile membrane filter 15-ml centrifugal filters, 5000 MWCO (e.g., Amicon filters; Millipore) Temperature-controlled centrifuge Lyophilizer or vacuum concentrator, optional Additional reagents and equipment for culture of hESC (see Section C in this book) 1. Remove and discard the standard hESC medium containing serum supplement and wash the hESCs with an equal volume of prewarmed (37◦ C) tissue-culture PBS. Repeat wash three more times. 2. Add the same volume of prewarmed (37◦ C), protein-free hESC media. Incubate cells according to standard hESC culture procedure for 24 hr. It is normal to see excess hESC shedding in the absence of any serum supplement.

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3. Collect the conditioned protein-free hESC medium and pass through a 0.22-µm sterile membrane filter. The conditioned medium can be frozen up to 2 years at −30◦ C (or lower) for later concentration and analysis.

4. Use centrifugal filtration at 4◦ C according to the manufacturer’s directions to concentrate the conditioned medium to less than 500 µl. 5. Wash once by adding 15 ml deionized water to 500 µl concentrate and using the same filter, reconcentrate to <500 µl to reduce the ionic strength. Using 15-ml, 5000 MWCO Amicon filters you can concentrate up to 50 ml of medium per filter by continually “topping up” the same filter.

6. Use the sample for tryptic digestion (Basic Protocol 5 or Alternate Protocol 5), analysis of protein concentration Basic Protocol 3 or Alternate Protocol 4), or gel electrophoresis (Basic Protocol 4) or store up to 2 years at −30◦ C. 7. Optional: To further increase protein concentration, reduce the volume or dry the extracellular protein extract using a lyophilizer or vacuum concentrator according to the manufacturer’s instructions. Dried samples can easily be reconstituted in 2-D gel extraction buffer, denaturing cell lysis buffer, or Laemmli buffer for SDS-PAGE analysis (see Reagents and Solutions). BASIC PROTOCOL 3

STANDARD PROTEIN QUANTIFICATION ASSAY This protocol is multilevel and is intended to quantify protein concentration in samples containing a variety of reagents. It is based on the original Bradford assay and is an adaptation of Bio-Rad’s protein microassay formatted for a 96-well flat-bottom plate. For information about the assay and a list of compatible reagents see the Bio-Rad protein assay technical manual (see Internet Resources). If the denaturing cell lysis buffer (Basic Protocol 1) was used without the addition of SDS, the sample can be diluted as little as 1.5-fold and assayed directly. If the 2-D gel-extraction procedure was used (Alternate Protocol 1) or if any reagents (e.g., see Table 1B.1.1) cannot be diluted below the acceptable limits have been used, a modified Bradford assay (see Alternate Protocol 4) will have to be performed. In the following format, this assay will accurately quantify protein in the range of 0.2 to 5 mg/ml. Samples above or below this range will have to be either diluted or concentrated (e.g., using a MWCO filter as in Basic Protocol 2 or by protein precipitation as in Support Protocol 1) to be assayed accurately.

Materials Protein sample (Basic Protocols 1 and 2; Alternate Protocols 1, 2, and 3) Sample diluent buffer (compatible with sample and assay reagents; e.g., see Internet Resources) 2 mg/ml (or greater) standard protein solution (BSA or IgG) Protein assay reagent (Bio-Rad) 96-well polystyrene flat-bottom microtiter plate 96-well microtiter plate reader with a 595-nm filter Graph paper or graphing program

Proteomic Analysis of Pluripotent Stem Cells

Prepare samples and standard 1. Select a volume of unknown protein sample to assay. Dilute to 80 µl with deionized water or appropriate compatible sample diluent buffer to get an approximate concentration of 1 mg/ml in order to maximize the number of data points available. NOTE: One must ensure that the concentration of buffer components after dilution (detergents, salts, other reagents) is compatible with the assay (see Table 1B.1.1).

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Table 1B.1.1 Compatibility of Common Reagents in Protein Extracts with the Indicated Assays

Bradford protein assay (Basic Protocol 3)a

2-D gel electrophoresisb (Basic Protocol 4)

Solution trypsin digestion (Alternate Protocol 5)

Urea

6M

8M

2M

Thiourea

1M

2 M with 7 M urea

NIc

Total combined detergent concentration should be <0.05%

0.25% (w/v) with >2% nonionic or zwitterionic detergents

0.05 % (w/v)

Triton X-100

4% (v/v) or 0.5% with 4% CHAPS

1 % (v/v)

CHAPS

4% (w/v)

1 % (w/v)

NP-40

4% (v/v)

1 % (v/v)

Tween 20

NIc

1 % (v/v)

Ionic salts buffers: <250 mM total

<15 mM total

<250 mM total

pH ∼ 2-9 (weak buffer)

pH ∼ 6-8

pH ∼ 8-9

DTT

100 mM

60 mM

20 mM

TCEP

Tested up to 10 mM

10 mM

5 mM

No limitd

No limite

No serine protease inhibitors f

Chaotropes

Detergents SDS

pH Reducing Agents

Protease Inhibitors a For a complete list of compatible reagents see the manual for the Bio-Rad protein assay reagent (see Internet Resources). b For more details about compatible reagents see the Amersham 2-D gel manual (see Internet Resources). c Abbreviation: NI, no information available. d Some protease inhibitors are amino acid based and can cause development of the protein assay reagent, resulting in artificially high

readings. Their effect on the background of the assay should be taken into account by including them in the dilution buffer for the assay at the same concentration as in the extracts. e An exception is some protease inhibitors that are (or contain) ionic salts (e.g., EDTA). The total salt concentration should not exceed 15 mM. f PMSF, a potent serine protease inhibitor has a half-life of ∼20 min in aqueous conditions. Protein extracts with fresh PMSF should be left on ice for at least 2 hr prior to digestion with trypsin.

2. Dilute 80 µg of protein standard (IgG or BSA) to a final volume of 80 µl in an appropriate buffer (one 80 µl standard per 96 well plate). The appropriate buffer will make the final concentration of reagents in the protein standard the same as the diluted unknown protein samples. For example, if assaying 20µl of samples in 0.1% (w/v) SDS and 8 M urea (diluting it in 60 µl of deionized water), the final concentrations in the 80 µl are 0.025% (w/v) SDS and 2 M urea. If the BSA protein standard was at 2 mg/ml in a weakly buffered solution (i.e., PBS), 40 µl of standard would have to be added to 40 µl of 0.05% (w/v) SDS and 4 M urea (sample diluent buffer) to make a final solution of 80 µg BSA in 80 µl 0.025% (w/v) SDS and 2 M urea.

3. Prepare a dilution buffer with components equivalent to the unknown protein sample and protein standard (∼1 ml per 96 well plate). If samples were prepared according to the above example, the dilution buffer would be 0.025% (w/v) SDS and 2 M urea.

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Figure 1B.1.1 Suggested layout for plating protein standards and diluted samples in the 96-well microtiter assay plate for the protein assay (Basic Protocol 3). All protein standards and samples are made up to 10 µl with assay diluent, and each well then has 200 µl of diluted protein assay reagent added per well.

Load and read plate 4. Dispense duplicate volumes of samples/standard into each row of the 96-well plate as outlined in Figure 1B.1.1. Fill each well with dilution buffer to a final volume of 10 µl. This will create a protein standard curve ranging from 0 to 10 mg of total protein as well as five different dilutions of unknown sample in duplicate.

5. Optional: To improve assay accuracy and account for light scattering from plate defects or bubbles in solution, take a background reading at a nonabsorbing wavelength (405 or 630 nm works for a Bradford assay) for each well before adding the reagent. 6. Dilute 1 part Bio-Rad protein assay reagent with four parts deionized water and dispense 200 µl into each well. Mix and allow 5 to 10 min at room temperature for development of the color. Protein-containing wells will shift from a red to a blue color. Pop bubbles with a toothpick or pipet tip to prevent interference with absorption reading.

7. Read absorbance at 595 nm for each well of the 96-well plate within 60 min on a microtiter plate reader.

Determine protein concentration 8. Subtract the background absorbance values (if taken in step 5) from the assay values at 595 nm for each well. The assay still has acceptable performance without background subtraction.

9. Average the duplicate values for each dilution of an unknown sample and each point of the protein standard. 10. Create a linear standard curve by plotting the averaged A595 values against the total amount of protein standard in each well. Using the layout prescribed here, 10, 8, 6, 4, 2, and 0 µl of diluted protein standard would correspond to 10, 8, 6, 4, 2, and 0 of total protein.

11. Optional: Ensure the linearity of the standard curve by checking its R2 value. If not >0.95 it may be necessary to disregard one or both of the 10 and 0 µg protein values. 12. Using only the A595 values that fall in the range of the corrected standard curve, extrapolate total protein values for each of the unknown sample dilutions. Proteomic Analysis of Pluripotent Stem Cells

13. Calculate the original concentration for each dilution by dividing the total protein detected by the undiluted volume plated in that well. For a given sample, average the concentration determined for all points which fall in the range of the standard curve.

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Protein concentration = (protein detected)/(volume of diluted sample) × (1/dilution) For example, assume 40 µl of original sample was diluted 1:2 to 80 µl. Then 5 µl of this dilution yielded 2.5 µg in the assay. The protein concentration of the original sample would be 1 µg/µl or 1 mg/ml.

MODIFIED PROTEIN QUANTIFICATION ASSAY This procedure is intended for protein samples with reagent concentrations that are incompatible with the standard protein assay. This assay is the same as the standard assay except all samples including the BSA or IgG protein standards are precipitated and reconstituted in a compatible buffer prior to assay.

ALTERNATE PROTOCOL 4

Additional Materials (also see Basic Protocol 3 and Support Protocol 1) Protein assay resolubilization buffer: 0.1% (w/v) SDS/8 M urea; prepared fresh or stored up to 1 year at −80◦ C Heating block set to 90◦ C 1. Using only deionized water as a diluent, prepare 80 µl dilutions of samples and protein standards as described in steps 1 and 2 of Basic Protocol 3. 2. Precipitate each unknown sample and protein standard according to Support Protocol 1. 3. Add 20 µl of protein assay resolubilization buffer to each protein precipitate. 4. Without disturbing the protein pellet, heat the pellet and protein assay resolubilization buffer 10 min at 90◦ C. After the pellet has gone into solution, dilute to 80 µl with deionized water. The concentration of reagents in the buffer will now be 2 M urea, 0.025% (w/v) SDS. This will also be the composition of the diluent buffer for dispensing samples into the microtiter plate for the assay (Fig. 1B.1.1 and Basic Protocol 3). Buffers containing urea should be prepared just prior to use, or frozen immediately at −80◦ C after preparation to avoid protein carbamylation which can interfere with proteomic analysis (McCarthy et al., 2003).

5. Centrifuge 2 min at 10,000 × g, room temperature, to ensure full resuspension of protein. 6. Use the reconstituted samples and protein standard to perform the standard protein assay as described in Basic Protocol 3, starting at step 3.

PRECIPITATION OF PROTEIN EXTRACTS The following procedure is a fairly universal method for precipitating proteins from solution. However, like all protein precipitation procedures, it is subject to a certain degree of sample loss. While the authors found this procedure very useful for most samples, there are a number of other procedures and kits (e.g., UPPA Protein Concentrate kit; Genotech) which may be an acceptable substitute.

SUPPORT PROTOCOL 1

Additional Materials (also see Basic Protocol 3) 2% (w/v) deoxycholate (DOC) solution 100% (w/v) trichloroacetic acid (TCA) solution Acetone, cooled to −20◦ C 1. Mix sample with 1/100 of its volume of 2% (w/v) DOC and incubate on ice for 30 min.

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2. Add enough 100% (w/v) TCA to bring the sample to a final TCA concentration of 15% (w/v) and vortex immediately for 30 sec to prevent any large precipitates from forming. 3. Let the sample stand and precipitate for a minimum of 1 hr on ice. If no precipitate is visible or time is an issue, samples can be left overnight at 4◦ C.

4. Collect precipitate by centrifuging 10 min at 15,000 × g, 4◦ C, and remove and discard the supernatant. Avoid disturbing the pellet. 5. Wash the pellet with 1 ml −20◦ C acetone and vortex to mix. The size of the pellet may be reduced as the DOC is washed out of the sample. If the pellet is compact and does not suspend, a 200-µl tip with the end cut off can be used to break apart the pellet by repeat pipetting.

6. Incubate the sample at least 10 min at −20◦ C (in the freezer). 7. Centrifuge samples 10 min at 15,000 × g, 4◦ C, and remove and discard the supernatant. Avoid disturbing the pellet. 8. Repeat steps 5 to 7. 9. Allow pellet to air dry to remove the acetone. Over-drying the pellet to the point where it becomes translucent can impede the resolubilization step. The protein precipitates need to be resolubilized in a buffer compatible with the next step of analysis. For example, use Laemmli buffer for 1-D gel analysis, extraction buffer for 2-D gel analysis (see Alternate Protocol 1), or a small volume of denaturing protein extraction buffer for a solution digest. If the protein precipitate is being used in the modified Bradford procedure, resuspend in resolubilization buffer as described in step 4 of Alternate Protocol 4. BASIC PROTOCOL 4

GEL ELECTROPHORESIS SEPARATION OF PROTEIN EXTRACTS The use of one-dimensional (1-D) gel electrophoresis, more commonly referred to as SDS-PAGE, is a common method for separating proteins based on molecular weight. Likewise, it is an excellent way to prefractionate protein samples prior to digestion and analysis by MS. A number of manufacturers sell all the necessary components required to run a 1-D gel in a variety of size formats. Moreover, there are many protocols available in generic scientific manuals (Sambrook et al., 1989; Gallagher, 2006). For the separation of complex proteomes and direct visual comparison of different samples, two-dimensional (2-D) gel electrophoresis has been the classic procedure of choice. In general, the most problematic aspect of 2-D gel electrophoresis is the isoelectric focusing (IEF) in the first dimension. It can be plagued with inconsistency and is subject to interference from ionic components in the sample. Use the recommended 2-D gel protein extraction protocol in Alternate Protocol 1 or see Table 1B.1.1 for a list of compatible reagents. The following protocol outlines a detailed generic protocol for successfully running the first dimension of this procedure. The main limitation of 2-D gel technology is that only the highest abundance proteins in a sample are visible. Consequently, to maximize sample loading and separation, running large format gels with 24-cm immobilized pH gradient (IPG) strips as outlined below, is recommended.

Materials Proteomic Analysis of Pluripotent Stem Cells

IPG strips (24-cm IPG Immobiline Drystrip pH 3 to 10 NL; GE Healthcare) Mineral oil Ampholytes, pH 3 to 10 (GE Healthcare)

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200 µg protein extract (Alternate Protocol 1) Two-dimensional (2-D) gel rehydration buffer (see recipe) containing protease inhibitors (see recipe) SDS equilibration buffer (see recipe) 1% (w/v) dithiothreitol (DTT) in SDS equilibration buffer (see recipe), freshly prepared 2.5% (w/v) iodoacetamide solution in SDS equilibration buffer (see recipe), freshly prepared Polyacrylamide slab gel (e.g., 25.5 cm × 20.5 cm × 0.15 cm; 10% or 12% (w/v) polyacrylamide recommended for whole-cell lysate) Protein molecular weight standards for electrophoresis, prestained, broad range (e.g., Bio-Rad) and recommended loading buffer, optional Agarose sealing solution (see recipe) SDS running buffer (see recipe) Trough for IPG strips Isoelectric focusing apparatus (e.g., Ettan IPGphor manifold and Ettan IPGphor; GE Healthcare) 25-ml serological pipets with ends cut off, optional Filter paper (absorbant electrophoresis type), optional Polyacrylamide gel electrophoresis apparatus, large format (minimum 25-cm wide), temperature control recommended (e.g., Bio-Rad) Microwave or hot water bath Run isoelectric focusing (IEF) gel 1. For each 2-D gel using 24-cm IPG strips, add the ampholytes according to the manufacturer’s instructions to 200 µg of protein extract and dilute to a final volume of 450 µl using 2-D gel rehydration buffer containing protease inhibitors. Ensure the ampholyte mixture matches the manufacturer and pH range of the IPG IEF strip.

2. Dispense protein solution evenly across the trough containing each 24-cm IPG IEF strip and cover with mineral oil to prevent evaporation. Allow strip to rehydrate 16 hr at room temperature. Do not perform this step below 15◦ C because the urea will come out of solution and poor rehydration will occur.

3. Focus the IPG strip in an Ettan IPGphor manifold with an Ettan IPGphor isoelectric focusing apparatus at the following voltage gradient (recommended as a starting point for standard hESC protein extracts):

1 hr constant 500 V 1 hr gradient to 1000 V 2 hr gradient to 5500 V 18 hr constant 5500 V. A total of just over 99 000 volt hr is generally required for complete isoelectric focusing of a 24-cm IPG strip. The progress of the IEF step can be gauged by the movement of the dye front in the IPG strip. Although protein focusing is far from complete when the bromphenol blue (BPB) from the rehydration buffer in the IPG strip has migrated to the anode, it is a good indication that interfering ionic species have been cleared from the center portion of the IPG strip. If the BPB has not migrated outside of the middle 2/3 of the IPG strip DO NOT apply a higher voltage. Instead, extend the 500 V step until the salts have been cleared. See manufacturers’ instructions for specific operation of focusing apparatus.

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4. Proceed to step 5 for reduction, alkylation, and equilibration in SDS equilibration buffer of the IPG strips, or freeze the IEF strips inside 25-ml serological pipets and store up to 1 month at −80◦ C. IEF strips can be frozen inside 25-ml serological pipets at −80◦ C and equilibrated at a later date without loss of resolution.

Reduce and alkylate IPG strips 5. To reduce the IPG strips, transfer the IPG IEF strips into freshly prepared 1% (w/v) dithiothreitol (DTT) in SDS equilibration buffer and incubate 15 min at room temperature, agitating periodically. Use enough solution to cover the entire strip. 6. To alkylate the free cysteine residues, transfer the IPG IEF strips into freshly prepared 2.5% (w/v) iodoacetamide solution in the SDS equilibration buffer and incubate 15 min at room temperature, agitating periodically. Use enough solution to cover the entire strip. Extending reduction and alkylation times beyond 15 min can result in loss of protein spot resolution. The return of the yellow colored acidic portion of the IPG strip to a deep blue is an indicator of complete reduction/alkylation.

Run SDS-PAGE 7. Transfer the reduced and alkylated IPG IEF strips to the top of the already cast polyacrylamide slab gels. Ensure that the gel portion of the IEF strip is in direct contact with the polyacrylamide gel surface, with the plastic backing against the glass, and that there are no air bubbles at the gel interface. Failing to ensure the plastic backing is against the glass can result in doubling or blurring of protein. A number of manufacturers offer systems to cast and run large-format polyacrylamide slab gels which are compatible with IEF strips. Regardless of the system used, all gels should be cast and ready to run at this point. Cold storage of focused IEF strips is an option, but at this point it is not recommended because it affects the quality of the resulting 2-D gel. For polyacrylamide slab gels to be compatible with the 24-cm IPG IEF strips they need to be at least 25 cm wide and 0.15 cm thick. A good starting point for a whole cell lysate is 10% or 12% (w/v) polyacrylamide. Gradient gels are not recommended because they make later 2-D gel comparisons very difficult.

8. Optional: Just adjacent to the end of each IPG IEF strip, place a square of filter paper (∼0.25 × 0.5 cm) soaked with electrophoresis protein molecular weight standards in the manufacturers’ recommended loading buffer. 9. Seal the strips (and standards) in place with the agarose sealing solution. Heat the solution briefly in either a microwave or boiling water bath to melt the agarose just prior to addition. 10. Run the gels (e.g., 12% (w/v) polyacrylamide slab gels, 25.5 × 20.5 × 0.15 cm) in SDS running buffer under the following conditions:

1 hr at 100 V (for stacking) 6 to 7 hr at 250 V (for completion) until the blue dye front migrates to within 0.5 cm of the bottom edge of the gel.

Proteomic Analysis of Pluripotent Stem Cells

To prevent warping and increase consistency the authors recommend holding the running buffer at a constant 15◦ C. See the manufacturers’ instructions prior to running the polyacrylamide gel apparatus.

11. Visualize protein spots as described in Support Protocol 2.

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COLLOIDAL COOMASSIE STAINING OF POLYACRYLAMIDE PROTEIN GELS

SUPPORT PROTOCOL 2

This staining procedure can be used for any protein gel where low background staining and high sensitivity are required. It is based on the previously reported Blue Silver staining protocol (Candiano et al., 2004). Other procedures (e.g., silver staining) are only slightly more sensitive than this method, but they can negatively impact downstream proteomic analysis. Fluorescent staining of protein gels can also be more sensitive; however the commercial reagents are costly and require specialized equipment for visualization and spot excision. Even though there are many acceptable staining alternatives, this procedure offers a very reproducible, sensitive, and inexpensive method for protein visualization and is well suited for comparative 2-D gel analysis. NOTE: In order to see optimal results from this staining protocol, one should allow 3 to 4 days for completion.

Materials Two-dimensional gel (2-D) gel with proteins (Basic Protocol 4) Protein gel fixing solution: 50% (v/v) ethanol 3% (v/v) phosphoric acid Modified Neuhoff’s solution (see recipe) Colloidal Coomassie staining solution: modified Neuhoff’s solution with 1.2 g/liter of Coomassie G250 Container(s) to hold gels Rotary shaker 1. Submerge the gel in protein gel fixing solution and gently shake for a minimum of 4 hr. Gels can be left in the fixing solution overnight or even for a few days. All volumes used in protocol steps should be sufficient to submerge the gels.

2. Drain and discard the excess fixing solution. 3. Wash the gel in modified Neuhoff’s solution 30 min with gentle shaking. Repeat twice more, changing the modified Neuhoff’s solution between each wash. 4. Drain the modified Neuhoff’s solution and cover the gel with colloidal Coomassie staining solution and shake for 3 to 4 days. Bright blue protein spots will be visible within the first 24 hr, but they can take 3 or 4 days to reach maximum intensity.

5. Following staining, store gel up to 1 year in modified Neuhoff’s solution (Coomassie free) in an airtight container.

IN-GEL TRYPSIN DIGESTION OF PROTEINS FOR IDENTIFICATION BY MASS SPECTROMETRY

BASIC PROTOCOL 5

This procedure has been designed for the digestion and extraction of proteins separated by polyacrylamide gel electrophoresis for the purpose of MS analysis. All volumes noted in this procedure have been optimized for spots from 2-D gels or small bands from 1D gels with a total volume of no more than 30 µl. Larger gel spots/bands require either division into multiple samples or a proportional increase in the volumes used. This procedure generally takes 3 days to complete with the steps broken down as follows: day 1, steps 1 to 3; day 2, steps 4 to 30; and day 3, steps 31 to 34. Characterization of Embryonic Stem Cells

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Materials Protein sample embedded in polyacrylamide gel (Basic Protocol 4) Acetonitrile/ammonium bicarbonate destaining solution: 1 M ammonium bicarbonate/20% (v/v) acetonitrile Methanol/acetic acid washing solution: 50% (v/v) methanol/5% (v/v) acetic acid Acetonitrile, HPLC grade 10 mM dithiothreitol (DTT), freshly prepared from 1 M stock solution (see recipe) Iodoacetamide 100 mM ammonium bicarbonate 50 mM ammonium bicarbonate, ice-cold Porcine modified trypsin, sequencing grade (Promega) 10% (v/v) formic acid in deionized (d)H2 O Scalpel 1.5- and 0.5-ml plastic microcentrifuge tubes Centrifuge 37◦ C water bath or incubator Vacuum concentrator (e.g., SpeedVac) NOTE: Anytime this protocol mentions the removal of a solution it will be necessary to centrifuge 30 sec at 5000 × g, room temperature, and aspirate the supernatant with a pipet.

Destain gel pieces 1. Excise the protein bands or spots from the gel as close to the stained area as possible, using a scalpel. Slice them into small pieces (∼1 mm to 2 mm square). While this is the most crucial step in optimizing sample recovery, crushing or pulverizing the gel pieces is not necessary because it will create fine particles that can later interfere with chromatographic analysis.

2. Place the gel pieces from each protein band or spot into a 1.5-ml plastic microcentrifuge tube. 3a. For gels stained with Coomassie (or a fluorescent stain like Sypro): Add 200 µl acetonitrile/ammonium bicarbonate destaining solution, vortex, and let stand for at least 4 hr at room temperature. Gel pieces can be left destaining overnight or for a few days.

3b. For silver-stained gels: See Support Protocol 3 for an alternate destaining procedure. 4. Remove and discard the destaining solution. 5. Add 200 µl methanol/acetic acid washing solution, vortex, and let the pieces stand 1 hr at room temperature. 6. Remove and discard the wash solution and add 200 µl acetonitrile/ammonium bicarbonate destaining solution. Let pieces stand 2 hr. 7. Repeat steps 4 to 6 until gel pieces are completely destained.

Reduce protein 8. Remove and discard the acetonitrile/ammonium bicarbonate washing solution.

Proteomic Analysis of Pluripotent Stem Cells

9. Dehydrate the gel pieces by adding 500 µl acetonitrile and, vortexing periodically, incubate 5 min at room temperature. When dehydrated, the gel pieces will have an opaque white color and will be significantly smaller in size.

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10. Remove the acetonitrile from the gel pieces and discard. 11. Allow the pieces to dry at room temperature. This process can be hastened by placing the gel pieces in a vacuum centrifuge for 2 to 3 min.

12. Prepare a fresh 10 mM DTT solution in 100 mM ammonium bicarbonate by combining 1.5 mg DTT per 1 ml of 100 mM ammonium bicarbonate solution. Vortex to dissolve. 13. Rehydrate the gel pieces with 30 µl freshly prepared 10 mM DTT solution and allow the pieces to reduce for 30 min at room temperature. 14. Ensure the gel pieces are fully rehydrated. Add more 10 mM DTT solution, if required. Gel pieces will swell and become transparent when rehydrated.

Alkylate protein 15. Remove and discard the DTT solution from the sample. 16. Prepare a fresh 100 mM iodoacetamide solution in 100 mM ammonium bicarbonate by combining 18 mg iodoacetamide per 1 ml of 100 mM ammonium bicarbonate. Vortex to dissolve. 17. Add 30 µl of 100 mM iodoacetamide solution and allow the pieces to alkylate for 30 min at room temperature. If an additional volume of 10 mM DTT was used in step 14, the same additional volume should be applied here.

18. Remove the iodoacetamide solution from the sample and discard. 19. Dehydrate gel pieces by adding 500 µl acetonitrile and, vortexing periodically, incubate 5 min at room temperature. 20. Remove the acetonitrile from the sample and discard. 21. Rehydrate the gel pieces in 200 µl of 100 mM ammonium bicarbonate. Vortexing periodically, let the reaction stand 10 min at room temperature.

Digest with trypsin 22. Remove any excess ammonium bicarbonate solution from the sample and discard. 23. Dehydrate gel pieces by adding 500 µl acetonitrile and vortex periodically for 5 min at room temperature. 24. Remove the acetonitrile from the sample and discard. 25. Completely dry the gel pieces in a vacuum centrifuge for 2 to 3 min at ambient temperature. 26. Prepare the fresh trypsin solution by adding 1 ml ice-cold 50 mM ammonium bicarbonate to 20 µg lyophilized trypsin (20 µg/ml final concentration). Keep the solution on ice until use. Any unused trypsin solution can be stored up to 1 year at −30◦ C, thawed on ice, and reused for later digests. Do not refreeze leftover solution.

27. Precool the samples on ice and add 30 µl trypsin solution to each. Allow the gel pieces to rehydrate on ice for 10 min, with occasional vortex mixing. Depending on the size, more than 30 µl may be required to fully rehydrate the gel pieces.

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28. Carefully remove the excess trypsin solution and discard. A small amount of trypsin, sufficient to digest the sample, is absorbed into the gel pieces during rehydration. The enzyme remaining in the gel after removal of excess solution will not begin to digest samples until they are incubated at 37◦ C in step 30.

29. Add 10 µl of 50 mM ammonium bicarbonate to the sample. Vortex and centrifuge the samples 30 sec at 5000 to 10,000 × g, 4◦ C. 30. Add enough 50 mM ammonium bicarbonate to completely cover the gel pieces and allow samples to digest overnight at 37◦ C.

Extract tryptic peptides 31. Add 30 µl of 50 mM ammonium bicarbonate to the digest and let stand 10 min at room temperature, with occasional gentle vortex mixing. 32. Centrifuge the samples 30 sec at 5000 to 10,000 × g, 4◦ C, and collect the supernatants into a separate 0.5-ml plastic microcentrifuge tubes. For this and the following steps 30 µl is the recommended default volume. Adjust this volume to cover all gel pieces with the extraction solutions.

33. Add 30 µl of 10% (v/v) formic acid solution to the gel pieces and let stand 10 min at room temperature, with occasional mixing. 34. Centrifuge the samples 30 sec at 5000 to 10,000 × g, 4◦ C, and combine the supernatant with the supernatant from step 32. 35. Repeat the steps 33 and 34 with a second 30-µl aliquot of 10% (v/v) formic acid solution, incubating 10 min. Extraction efficiencies can be slightly improved by performing extraction incubations in a sonicating water bath.

36. Perform a final extraction by adding 200 µl acetonitrile to the gel pieces. Allow the pieces to dehydrate 10 min, with occasional vortex mixing. 37. Centrifuge the samples 30 sec at 5000 to 10,000 × g, 4◦ C, and combine the supernatant with the extracts from steps 32 and 34. 38. For MS analysis, reduce the volume of the peptide extracts to less 20 µl by evaporation in a vacuum centrifuge at ambient temperature. Do not allow the extract to dry completely. If the extracts do dry completely, reconstitute with 20 µl of 10% (v/v) formic acid solution. SUPPORT PROTOCOL 3

DESTAINING SILVER-STAINED GELS Silver staining is not a preferred method of detection for protein gels destined for MS analysis. However, if the polyacrylamide protein gel has been silver stained, an alternative destaining protocol is necessary prior to trypsin digestion.

Materials Silver-stained sectioned gel pieces (Basic Protocol 5, steps 1 and 2) 30 mM potassium ferricyanide 100 mM sodium thiosulfate Acetonitrile/ammonium bicarbonate destaining solution: 1 M ammonium bicarbonate/20% w/v acetronitrile Proteomic Analysis of Pluripotent Stem Cells

1. Just prior to use, mix 30 mM potassium ferricyanide and 100 mM sodium thiosulfate at a 1:1 (v/v) ratio to make 500 µl/gel working destaining solution.

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2. Cover the silver-stained pieces with the acetonitrile/ammonium bicarbonate destaining solution and incubate at room temperature, with periodic vortex mixing until the brown color disappears from the gel pieces (normally 5 to 30 min). 3. Remove the destaining solution by centrifuging 30 sec at 5000 × g, room temperature, and aspirating the supernatant with a pipet. Rinse the gel pieces twice with 0.5 ml deionized water, centrifuging and aspirating the supernatant between rinses. 4. Remove and discard the deionized water and cover the gel pieces with 200 µl acetonitrile/ammonium bicarbonate destaining solution. Let stand at least 1 hr at room temperature with occasional vortex mixing. Gel pieces can be left in the destaining solution overnight or even for a few days.

5. Proceed with trypsinization and analysis, starting at step 8 in Basic Protocol 5.

IN-SOLUTION TRYPSIN DIGESTION OF PROTEINS FOR IDENTIFICATION BY MASS SPECTROMETRY

ALTERNATE PROTOCOL 5

This protocol outlines a procedure for digesting proteins still in solution with trypsin for MS analysis. Lower sample volumes and higher protein concentrations maximize digestion efficiency. Trypsin is subject to interference with a number of reagents common in protein extracts. Please refer to Table 1B.1.1 for a detailed list of reagent concentrations compatible with the trypsin digest. Because many of these reagents (especially detergents) can be difficult to remove post-digestion, it may be desirable to remove them prior to digestion. Read the entire protocol in advance and formulate a strategy to digest the protein and obtain peptides that can be analyzed by MS.

Materials Protein sample (from Basic Protocol 1 or 2 or Alternate Protocol 1, 2, or 3, extracted using Support Protocol 1) Denaturing cell lysis buffer (see recipe) 50 mM ammonium bicarbonate in dH2 O Dithiothreitol (DTT), solid or 1 M stock solution (see recipe) 100 mM ammonium bicarbonate in deionized (d)H2 O Iodoacetamide, solid Porcine modified trypsin, sequencing grade (Promega) Acetonitrile, HPLC grade Heating block, set at 65◦ C (optional) 37◦ C water bath or incubator Prepare sample 1a. If the sample is dry, pelleted, or lyophilized: Suspend in 50 mM ammonium bicarbonate at a protein concentration of ∼1 mg/ml. 1b. If the sample solubility is questionable: Employ Basic Protocol 1, step 2b and treat the protein pellet as if it were a cell pellet. If not using one of the standard extraction protocols recommended, ensure the pH of the protein extract is between 8 and 9 prior to the reduction step.

Reduce protein 2. Make up a fresh 200 mM DTT stock by adding 30 mg DTT solid to 1 ml of 100 mM ammonium bicarbonate. Alternatively, the DTT solution can be made by diluting 1 mM DTT stock (see recipe) 1:5 with 100 mM ammonium bicarbonate.

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3. Reduce the sample by adding 5 µl of 200 mM DTT stock to 100 µl sample. Let stand at ambient temperature for 45 min, mixing periodically by vortexing. 4. Alternatively (if the sample does not contain urea), add 50 µl of 200 mM DTT per 100 µl sample, and heat 10 min at 65◦ C to reduce the proteins. CAUTION: Without the presence of chaotropes (e.g., urea) or detergents (e.g., SDS or CHAPS) protein can easily precipitate under reducing conditions in the presence of heat.

5. Centrifuge 30 sec at 5000 × g, 4◦ C, to collect the sample in the bottom of the tube.

Alkylate protein 6. Make up a fresh stock solution of iodoacetamide by adding 36 mg of iodoacetamide solid to 1 ml of 100 mM ammonium bicarbonate (200 mM iodoacetamide final concentration). 7. Alkylate the sample by adding 20 µl of the 200 mM iodoacetamide stock per 100 µl of sample. Let stand at ambient temperature for 45 min and vortex periodically. 8. Neutralize the remaining iodoacetamide by adding 10 µl of 200 mM DTT stock solution per 100 µl of sample. Let stand at ambient temperature for 45 min and vortex periodically.

Digest with trypsin 9. Dilute sample with 50 mM ammonium bicarbonate so that all of its buffer components are within compatible concentration ranges (see Table 1B.1.1). Remember to also factor in the volume of trypsin to be added in step 10. 10. Determine the amount of trypsin required for the digest and prepare it from the lyophilized stock using the 50 mM ammonium bicarbonate solution. Reconstituted trypsin can be freshly frozen once at < −30◦ C and reused. Generally, the ratio of trypsin to sample should be at least 1:25 for a successful digest meaning 25 µg of protein in a sample requires 1 µg of trypsin to be digested properly in solution. If the total amount of protein in a sample is unknown try staring with 10 µg for any digest with a volume under 1 ml. For example, assume the original protein concentration was 1 mg/ml in 8 M urea and 50 mM ammonium bicarbonate and you were digesting 100 µl (100 µg protein) of sample. The volume after reduction and alkylation would be 135 µl. Consequently, one would need to add 4 µg of trypsin in a total of 400 µl of 50 mM ammonium bicarbonate to properly digest the sample while keeping the concentration of urea compatible at <1.5 M.

11. Briefly vortex sample to mix. Allow the sample to digest overnight at 37◦ C (or for at least 18 hr).

Clean up digest 12a. To remove detergents (e.g., Tween, NP-40, CHAPS, Triton-X, or SDS) from sample: Use protein precipitation prior to digestion (Support Protocol 1) for samples containing proteins or strong cation exchange following digestion (SCX; Basic Protocol 7) for samples that contain peptides. 12b. To remove interfering protease inhibitors (along with detergents) from sample preparation: Precipitate proteins (see Support Protocol 1) prior to trypsin digestion (steps 9 to 11). Proteomic Analysis of Pluripotent Stem Cells

13. Using sample with interfering substances removed (if necessary), proceed to Basic Protocol 6 to concentrate and desalt peptides prior to analysis by mass spectrometry.

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The trypsin (protein) in the solution can potentially interfere with identification of peptides by MS. However, the concentrations specified and the use of modified trypsin minimize its masking effect (see Troubleshooting for more details). Moreover, during cleanup (Basic Protocol 6), a majority of the trypsin is retained on the C18 cartridge and not eluted with the digested peptides.

DESALTING AND CONCENTRATION OF PROTEINS/PEPTIDES BY REVERSE-PHASE CHROMATOGRAPHY

BASIC PROTOCOL 6

This procedure, based on hydrophobic interactions, facilitates concentration of proteins/peptides while allowing the removal of hydrophilic buffer components prior to MS analysis. Detergents, which will interfere with subsequent MS analysis, will also be copurified by this method and, consequently, must first be removed. To remove detergents use protein precipitation (Support Protocol 1) for samples containing proteins or strong cation exchange (Basic Protocol 7) for samples that contain peptides.

Materials Buffer B: 0.1% (v/v) formic acid in acetonitrile Buffer A: 0.1% (v/v) formic acid in deionized H2 O Sample (Alternate Protocol 5) 10% (v/v) formic acid Buffer C: mix Buffer A and Buffer B 1:1 Disposable C18 solid phase extraction (SPE) column, 1 ml capacity ∼100 mg solid phase Vacuum apparatus/manifold 1.5- and 0.5-ml microcentrifuge tube Vacuum centrifuge (e.g., SpeedVac) 1. Activate and clean the C18 SPE cartridge by passing 1 ml of buffer B through it, discarding the effluent. Repeat three times. To maximize efficiency of the following procedure ensure that once wet, the solid phase resin of the C18 SPE does not dry out or have air passed through it. Use the vacuum manifold or a pipet to force samples and buffers through the SPE cartridge, or according to manufacturers’ recommendations.

2. Prepare the C18 SPE cartridge for binding by passing 1 ml of buffer A through it, discarding the effluent. Repeat three times. 3. Acidify the sample by diluting it 1:2 with 10% (v/v) formic acid in deionized water. CAUTION: Carbonate-based buffers may foam slightly. Use a large enough container to avoid sample loss.

4. Pass the sample over the SPE cartridge at a speed no faster than 1 drop per second, discarding the effluent. 5. Wash away hydrophilic buffer components from the sample by passing 1 ml of buffer A over the SPE cartridge, discarding the effluent. Repeat 3 times. 6. Elute the peptides from the column with 400 µl buffer C at a speed no faster than 1 drop per second. Collect the effluent in a microcentrifuge tube. 7. Optional: To elute larger compounds (proteins) repeat the elution with 400 µl of buffer B. 8. Reduce the sample volume and remove the acetonitrile in a vacuum centrifuge as described in Basic Protocol 5, step 38. If sample is brought to dryness it can be reconstituted in a small volume of 10% (v/v) formic acid prior to MS analysis. Current Protocols in Stem Cell Biology

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BASIC PROTOCOL 7

STRONG CATION EXCHANGE (SCX) EXTRACTION AND FRACTIONATION OF SAMPLES This procedure, based on interactions with a strongly acidic solid phase, facilitates concentration of proteins or peptides while allowing the removal of neutral and anionic buffer components (most detergents) prior to MS analysis. Using a single-step elution, interfering substances can be removed; however, by utilizing a stepwise elution method, complex tryptic digests can be prefractionated prior to MS analysis to greatly increase depth of analysis. The later application is more commonly referred to as multidimensional protein identification technology (MuD-PIT; Washburn et al., 2001; Delahunty and Yates, 2005; Chen et al., 2006). The following procedure has been optimized using a basic highpressure liquid chromatography (HPLC) set-up, however it could be easily adapted to hand-held SCX cartridges. NOTE: This procedure is subject to sample loss and is not recommended for samples containing <25 µg of protein. Generally SCX resin, based on a sulfonic acid support, can retain approximately 15 mg of protein per ml of packing.

Materials Mobile phase: 0.1% (v/v) formic acid, 20% (v/v) acetonitrile in H2 O 50 to 500 µg protein sample: total salt concentration (i.e., Na+ , K+ , NH4 + ) <10 mM; diluted with mobile phase or desalted as in Basic Protocol 6. KCl stock: 500 mM potassium chloride (KCl)/20% (v/v) acetonitrile/0.1% (v/v) formic acid 0.8 × 50–mm cation exchange column (Bio SCX Series II; Agilent) Analytical HPLC equipment: flow rate of 1 ml/min, minimum 20-µl sample loop Vacuum centrifuge (e.g., SpeedVac) Prepare columns 1. Prepare and wash SCX column according to manufacturer’s instructions. 2. Equilibrate SCX column in mobile phase (5 ml at 0.5 to 1 ml per min of mobile phase, for this example).

Prepare samples 3a. For samples with <10 mM total ionic salts: Acidify using neat formic acid to pH 2 to 3.5. 3b. For samples with high salt concentration: Dilute with SCX mobile phase to a salt concentration of <10 mM and acidify to pH 2 to 3.5. 4. Load at least 50 µg (50 to 500 µg for the column in this example) of protein or peptide digest onto the column. Some peptides will not stick to the column and will instead wash through. If performing a MuD-PIT experiment (see protocol introduction) and the sample is free of detergents, collect the flow through as the 0 mM fraction for later MS analysis.

5. Wash the sample on column with mobile phase (5 ml for this example).

Run columns If performing a single step clean-up 6a. Elute digested peptides with 40 µl of 300 mM KCl. Proteomic Analysis of Pluripotent Stem Cells

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Whole or undigested proteins (e.g., trypsin) may be eluted by repeating this step but injecting 40 µl of 500 mM KCl stock solution.

7a. Taking into account the HPLC dead volume, collect the first 1 ml of the elution into a 1.5-ml microcentrifuge tube. Current Protocols in Stem Cell Biology

If performing a MuD-PIT experiment 6b. Make up 1 ml of each of the following KCl solutions by mixing the 500 mM KCl stock and mobile phase: 7.5, 15, 30, 45, 60, 75, 90, 120, 150, 300, and 500 mM KCl. Elute the first fraction by injecting 20 µl of the 7.5 mM KCl solution. 7b. Taking into account the HPLC dead volume, collect the first 1 ml of the elution into a microcentrifuge tube. Repeat this step for each increasing KCl elution buffer from 15 mM to 500 mM. This step prefractionates a complex peptide mixture for MS analysis.

8. Bring fractions to dryness in a vacuum centrifuge. 9. Reconstitute samples in a small volume of 10% (v/v) formic acid and desalt according to Basic Protocol 6 or, preferably, during reversed-phase LC-MS/MS analysis (see Steen and Mann, 2004).

REAGENTS AND SOLUTIONS Unless otherwise stated all solutions should be prepared in the highest quality deionized water. Either 18 mega-ohm d H2 O passed through at 0.22-µm filter or commercial HPLC-grade deionized water is sufficient. All other reagents should be of the highest quality possible, electrophoresis grade or better. A number of solutions (e.g., SDS, DTT, EDTA, and Tris·Cl) can be prepared ahead of time to ease preparation of other buffers.

Affinity extraction buffer 50 mM Tris·Cl, pH 7.4 (see recipe) 150 mM NaCl 1% (v/v) NP-40 or Triton X-100 Store up to 1 year at −30◦ C or 1 week at 4◦ C Agarose sealing solution 0.5% (w/v) agarose 0.1% (w/v) SDS: add using 10% (w/v) SDS stock solution (see recipe) 0.002% (w/v) bromphenol blue Store up to 1 year at 4◦ C Heat before use to liquefy agarose The solution will form a gel-like solid at room temperature.

Denaturing cell lysis buffer 20 mM ammonium bicarbonate 8 M urea 2% (w/v) SDS: add using 10% (w/v) stock solution (see recipe) Prepare fresh just prior to use Dithiothreitol (DTT) stock solution, 1 M Add 1.55 g of DTT to 5 ml water. Once dissolved, dispense into 250-µl aliquots, and immediately store up to 1 year at −30◦ C. Do not refreeze thawed aliquots more than two times.

EDTA, 500 mM Dissolve 18.7 g of EDTA in 80 ml of water. Raise pH with concentrated NaOH or ∼2 g of solid NaOH. Heat and dissolve with mixing. Once dissolved adjust pH to 8.0 with concentrated HCl and make up to 100 ml with water. Store up to 1 year at room temperature.

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Hypotonic cell lysis buffer 20 mM Tris·Cl, pH 7.4 (see recipe) 10 mM NaCl 5 mM MgCl2 5 mM CaCl2 1 mM DTT (add fresh from 1M stock; see recipe) Store up to 1 year at −30◦ C Prepare fresh This buffer can also be prepared without DTT and frozen up to 1 year at −30◦ C, with the DTT added just prior to use.

Laemmli buffer, 1× 2% (w/v): add using 10% (w/v) stock solution (see recipe) 10% (v/v) glycerol 50 mM Tris·Cl, pH 6.8 (see recipe) 0.01% (w/v) bromphenol blue 100 mM dithiothreitol (DTT) or 5% (v/v) 2-mercaptoethanol This is usually prepared as a 2× to 5× stock solution and stored at < −20◦ C in small aliquots to minimize freeze-thaw cycles. The SDS in Laemmli buffer will precipitate when cold. Before use, warm the Laemmli buffer to ambient temperature, add the buffer to the cells, and immediately heat to 65◦ C to prevent sample degradation.

Modified Neuhoff’s solution 10% (w/v) ammonium sulfate 20% (v/v) methanol 10% (v/v) phosphoric acid Prepare fresh Nuclease cocktail, 20× stock 1 mg/ml DNase I (GE Healthcare) 0.25 mg/ml RNase A (Sigma) 50 mM MgCl2 5% (v/v) glycerol Once in solution, divide into 100-µl aliquots and store up to 1 year at −20◦ C. Dilute to a 1× concentration in cell lysate and incubate at ambient temperature up to 30 min until viscosity of the lysate is reduced. Nucleases are inhibited by the presence of metal chelators such as EDTA. The MgCl2 can interfere with IEF in 2-D gel experimentation, and it also inhibits nucleases, so use accordingly.

Phosphatase inhibitor cocktail 50× sodium fluoride: 1 M sodium fluoride in water, store up to 1 year at −20◦ C 50× 2-glycerolphosphate: 1 M β-glycerolphosphate in water, store up to 1 year at −20◦ C

Proteomic Analysis of Pluripotent Stem Cells

200× sodium orthovanadate: Add 0.37 g to 9 ml water and adjust pH to 10 with concentrated HCl or NaOH (solution will become brown/yellow). Heat to 100◦ C until solution turns colorless (∼10 min). Cool to room temperature and readjust pH to 10. Repeat until solution remains colorless and pH stabilizes at 10. Bring volume to 10 ml with water, divide into 100-µl aliquots, and store up to 1 year at −20◦ C. continued

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Dilute each inhibitor to 1× concentration in the cell lysis buffer being used for the experiment and use just prior to performing cell lysis. The high ionic strength of these reagents can interfere with IEF in 2-D gel experimentation, so use accordingly. Each of these inhibitor stocks should be prepared individually and stored as described. Store aliquots in small volumes to avoid multiple freeze-thaw cycles. A number of manufactures offer premixed phosphatase inhibitor cocktails which are acceptable. The above recipe is for a generic phosphatase inhibitor cocktail that is commonly employed in mammalian cell studies.

Protease inhibitor cocktail 100× phenylmethylsulphonyl fluoride (PMSF; 100 mM): prepared in isopropanol and stored in 100-µl aliquots up to 1 year at −20◦ C 100× leupeptin (2 mM): prepared in water and stored in 100-µl aliquots up to 1 year at −20◦ C 100× pepstatin (150 µM): prepared in methanol and stored in 100-µl aliquots up to 1 year at −20◦ C 500× EDTA (500 mM; see recipe) Dilute each inhibitor to 1× concentration in the cell lysis buffer being used for the experiment and use just prior to performing cell lysis. The high ionic strength of EDTA can interfere with IEF in 2-D gel experimentation and also inhibits nucleases, so use accordingly. Each of these inhibitor stocks should be prepared individually and stored as described. Store aliquot in small volumes to avoid multiple freeze-thaw cycles. Together these reagents will inhibit serine proteases (PMSF and leupeptin), cysteine proteases (leupeptin), aspartic acid proteases (pepstatin), and metalloproteases (EDTA). A number of manufactures offer premixed protease inhibitor cocktails which are acceptable. The above recipe is for a generic protease inhibitor cocktail that covers the spectrum of proteases common in mammalian cells.

Protein-free hESC medium High-glucose DMEM 1% (v/v) 100× MEM nonessential amino acids solution (Gibco/Invitrogen) 1 mM L-glutamine 0.1 mM 2-mercaptoethanol Sterilize by passing through a 0.22-µm filter under vacuum. Store up to 1 year at −30◦ C or 1 month at 4◦ C. Just prior to use add protein growth factors (e.g., 8 ng/ml basic fibroblast growth factor; Invitrogen), if desired. SDS equilibration buffer 2% (w/v) SDS: add using 10% (w/v) stock solution (see recipe) 6 M urea 30% (v/v) glycerol 50 mM Tris·Cl, pH 8.0 (see recipe) Prepare just prior to use or store up to 1 year at −80◦ C. Buffers containing urea should be prepared just prior to use, or frozen immediately at −80◦ C after preparation to avoid protein carbamylation, which can interfere with proteomic analysis (McCarthy et al., 2003).

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SDS running buffer, 10× stock 121.1.g Tris base 576 g glycine pH 8.3 40 g SDS H2 O to 4 liters Store up to 1 year at room temperature Before using, dilute to 1x with water (final concentrations 25 mM Tris, 192 mM glycine, 0.1% SDS).

SDS stock solution, 10% (w/v) Add 5 g of SDS in 45 ml of water Mix and heat to dissolve. Once SDS is in solution make up to 50 ml with water. Store indefinitely at room temperature. SDS can be particularly insoluble and hard to work with. This 10% solution is meant to aid in the preparation of other SDS-containing buffers, namely SDS equilibration buffer, Laemmli buffer, agarose sealing solution, and the protein assay resolubilization buffer in Alternate Protocol 4.

Tris·Cl (1 M), pH 7.4 To make 1 liter: Dissolve 121.1 g of Tris base in 800 ml of water. Adjust pH to 7.4 with concentrated HCl (∼70 ml). Wait until solution cools to make final pH adjustment. Dilute to 1 liter with water. Store up to 1 year at room temperature. To make solutions of different molarities and pHs adjust the amounts of Tris base and HCl used.

Two-dimensional gel extraction buffer 4% (w/v) CHAPS 20 mM ammonium bicarbonate 20 mM DTT 7 M urea 2 M thiourea Store up to 1 year at −80◦ C. Buffers containing urea should be prepared just prior to use or frozen immediately at −80◦ C after preparation to avoid protein carbamylation which can interfere with proteomic analysis (McCarthy et al., 2003).

Two-dimensional gel rehydration buffer 6 M urea 2 M thiourea 4% (w/v) CHAPS 0.4% (w/v) DTT 0.5% (v/v) ampholytes (GE Healthcare) 10 µl/ml Ettan protease inhibitor mix (GE Healthcare) 1× (1 mM) PMSF (use 100× solution; see recipe for protease inhibitor cocktail) ∼0.0001% (w/v) of bromphenol blue Make up the solution containing urea, thiourea, CHAPS, DTT, and bromphenol blue at the indicated final concentrations in deionized water and store in 10-ml aliquots up to 1 year at −80◦ C. Just prior to use, thaw at ambient temperature and add protease inhibitor mix, PMSF, and ampholytes. Proteomic Analysis of Pluripotent Stem Cells

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COMMENTARY Background Information Identification of proteins in complex mixtures The biggest difference between mass spectrometry-based proteomics and many other high-throughput, array-based technologies is the predictable outcome of the experiment. In array-based screens, the candidates on the array are already known and consequently, so are the potential outcomes. In contrast, mass spectrometry-based proteomic screens are true “discovery-based” approaches, in that the analysis is de novo, meaning the possible outcome of the experiment cannot be predicted from the beginning. Another challenging aspect of using mass spectrometry is this: even if a protein of interest is not detected, it may still be present in a sample. Experimental conditions and the time allowed for the analysis may not have been optimal for detecting certain proteins, especially when looking for lower-abundance proteins in complex mixtures. Consequently, if it is already known which protein is being sought, and it can be detected by some other means (e.g., western blotting), then using that technology may be the more sensitive, efficient, and cost-effective approach. Searching for proteins that are at relatively low concentrations or have been posttranslationally modified (e.g., phosphorylated) in complex samples (e.g., cell lysate) presents a substantial hurdle. Quite often, species present at higher concentrations tend to eclipse the signals of the less abundant proteins during mass spectrometry–based proteomic analyses. To combat this drawback, much effort has been put into more advanced instrumentation as well as better sample preparation prior to analysis. One approach that has been developed to increase the chance of identifying more proteins and even lower-abundance proteins in a complex mixture involves sample fractionation. Several of these strategies are described in this unit: subcellular fractionation (Alternate Protocol 3), one-dimensional SDS-PAGE separation (Basic Protocol 4), and MuD-PIT separation of tryptic peptides (Basic Protocol 7). When the analyses are focused upon identifying protein complexes or posttranslational modifications, affinity selection methods are the best choice to separate these species away from their higher-abundance counterparts prior to analysis. To find a more

in-depth review of these strategies, see Key References at the end of this unit. Quantitative proteomics Perhaps the most widely used and earliest developed method to look at changes at the protein level is analysis by two-dimensional (2-D) gel electrophoresis. In this method quantitation of proteins between different samples is achieved by the software-based comparison of gel images, which allows the identification of differentially expressed proteins. While it is not the most modern tool used to examine differences at the proteome level, 2-D gel electrophoresis has several distinct advantages, including speed of operation, high sample loading capacity, and cost effectiveness. Another benefit is that proteins of interest can be selected for mass spectrometric analysis by sampling only from identified spots on the gel, without the need to analyze all of proteins in the sample. Unfortunately, 2-D gel electrophoresis also carries with it several disadvantages. For instance, protein solubility can be an issue, because membrane and hydrophobic proteins can precipitate during isoelectric focusing. This can lead to decreased protein spot resolution and streaking of the gel images. In addition, the isoelectric focusing step is very sensitive to interference from ionic compounds like salts, nucleic acids, and lipids present in the sample. While 2-D gels allow for high sample loading, only the most abundant proteins will ever be visible on the gel for analysis. Consequently, even though 2-D gel electrophoresis is a good first step in the comparison of different proteomes, sample prefractionation or other quantitative proteomic techniques will be necessary to study the lower-abundance proteins in a sample. Newer approaches for quantitatively evaluating complex proteomes rely on comparing the signal (number of counts) obtained by mass spectrometry for a given protein. The majority of these techniques use stable isotopes (e.g., 12 C versus 13 C) to differentiate two or more samples. The stable isotopes are incorporated into each of the samples and then the samples are mixed. Relative quantitation is achieved by comparing the mass spectral signals from the “heavy” (e.g., sample labeled with 13 C) and “light” (i.e., sample labeled with 12 C) labeled samples. The majority of isotopic incorporation is achieved by exploiting amino acid

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functional groups such as sulfhydryl, amine, carboxy, or carbohydrate attachment sites. Alternatively, isotopically coded amino acids can be added directly into cell culture growth media, allowing the target cells to accumulate isotopically labeled amino acids that are later identified by mass spectrometric analysis. This method is termed stable isotope labeling with amino acids in culture (SILAC; Ong et al., 2002; Mann, 2006), and it avoids problems (e.g., increased sample handling) experienced during typical chemical modifications of samples following protein extraction. Currently, while no one isotopic incorporation strategy has come to the forefront in proteomics, their popularity continues to increase as noted by Ong and Mann (2005) in a recent comprehensive review. The biggest drawback to these isotopebased technologies is the high cost of the specialized isotopic reagents. Other disadvantages include the limited availability of software for data analysis, as well as the heavy time demand on mass spectrometry instrumentation for these comprehensive experiments. Such factors can make these approaches costprohibitive or inaccessible to the average biological science laboratory.

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Mass spectrometric analyses Just as there is a wide range of proteomic methodologies, there are many different types of mass spectrometers available, each with its own advantages and disadvantages (Glish and Vachet, 2003). The simplest and fastest type of mass spectrometer commonly employed in proteomic analysis is a matrix-assisted laserdesorption ionization time-of-flight (MALDITOF) mass spectrometer. While this instrument produces mass spectral data very quickly, it lacks the ability to fragment peptide ions and hence, cannot determine individual peptide sequences. Consequently, it is best utilized to identify digests of single proteins and is most often used to identify proteins from a spot from a 2-D gel. For more complex protein digests or for spots containing more than one component, the most commonly employed instruments are electrospray ionization (ESI) coupled with ion traps and quadrupole time-of-flight (Q-TOF) instruments. Other instruments with increasing popularity despite their high costs are the Fouriertransform ion-cyclotron-resonance mass spectrometer (FT-ICR MS) and the Orbitrap. These provide much higher resolution and mass accuracy but often at the expense of sensitivity. An advantage of these electrospray ionization

instruments is that they can be easily interfaced to an HPLC to provide separation of tryptic peptides during analysis. The eluting peptides are ionized by electrospray ionization (ESI). Moreover, as each peptide is eluted into the instrument, it can be isolated and fragmented in order to determine its amino acid sequence. While ion trap-based instruments tend to be less expensive and more robust than Q-TOFs, their mass accuracy and effective mass range are significantly lower. As a result, they can generate a lot more false positive protein identifications. Regardless of the type of MS instrument, it is critical that they are frequently calibrated for mass accuracy and resolution. As technology advances in the field of mass spectrometry, new instruments are constantly being developed that serve to separate and identify proteins, peptides, or small molecules. For example the latest generation of nano liquid chromatography instruments (nano LC) provide much more accurate and reproducible retention times such that these can be reliably used as parameters in MS experiments (for exclusion list) or even for peptide identification (Krokhin et al., 2004). A review encompassing a more complete discussion of MS instrumentation basics, their proteomic applications, and sequencing of peptides by MS-based fragmentation can be found in the Key References at the end of this unit. Interpretation of mass spectrometry data Interpretation of MS data can range from the very simple (e.g., identifying the molecular weight of a single protein) to the very tedious (e.g., large scale quantitative or PTM analysis of a complex mixture) and is often the most time-consuming aspects of a proteomic experiment. Fortunately, there are a number of software packages available from MS manufacturers or from third-party providers (e.g., PEAKS, BSI; MASCOT, Matrix Science; X!, the GPM; see Internet Resources) for the interpretation of mass spectral data, including protein identification, post-translational modifications and quantitation. However, the compatibility of the different available software packages with data format varies both with instrument types and manufacturers. While attempts have been made to make these platforms more uniform (Pedrioli et al., 2004), much remains to be done to have data formats that are truly instrument independent. Our recommendation is to consult with the MS service provider about the desired experimental outcome and get their input on the software that might be most effective for reaching one’s

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goals. The authors’ experience indicates that it is worthwhile to use more than one software and database to increase confidence in protein identification. For more details and online proteomics tools, please refer to Internet Resources at the end of this unit.

Critical Parameters and Troubleshooting The primary problem that arises in a general proteomic analysis/identification is that the proteins of interest are not identified. Instead, proteins that should not be present in the sample are identified as the primary candidates (or “top hits” in a database search). A list of the most commonly identified (and unwanted) proteins with solutions for getting rid of them includes: Serum albumin. This is the most abundant protein in serum replacement media used to culture hESCs. Continual identification of this protein, can usually be attributed to insufficient washing of the cells prior to lysis. Solution: Ensure the cells are washed thoroughly and often with PBS just prior to extraction. Transferrin. This is the second most abundant protein in serum replacement media used to culture hESCs. Solution: Apply the same precautions as used for serum albumin. Keratin. In general, this is the most common contaminant in proteomic samples. Keratin comes directly from people handling the samples and shedding skin cells, as well as from dust in the air and on laboratory surfaces. Refer to the cautionary notes in the introduction of this unit on tips for keeping samples keratin-free. Immunoglobulin (Ig). While also a major serum protein, Ig typically appears in the list of identified proteins in experiments that have employed affinity applications using antibodies. Whether purifying protein complexes or phosphorylated proteins, Ig can end up in the sample when the compound of interest is eluted from its solid support. Solutions: (1) Perform SDS-PAGE on the sample prior to MS analysis with Ig in a control lane so you are aware of which bands correspond to the Ig heavy and light chains. (2) Instead of using protein A/G to bind Ig to its support, covalently crosslink it using amine or sulfhydryl reactive media. Trypsin. This is the one protein will be difficult (if not impossible) to remove from sample because it is required to generate peptides for MS analysis. Solution: The trypsin digest protocols described here (Basic Protocol 5 and

Alternate Protocol 5) have been optimized to minimize the presence of trypsin peptides in the sample. However, if trypsin is the primary protein identified (i.e., the “top hit” in a database search), feel reassured that the digest and analysis were performed properly. Unfortunately, it also means that there was very little, if any, other protein in the digested sample. The next most common problem is that one identifies a number of proteins, but sees nothing of “biological interest.” Unfortunately, biologically interesting proteins tend to be of lower abundance and are generally not detected in a quick analysis of a complex mixture. There is no definitive solution for this problem, and it remains an intensive area of proteomic method development. To date, the best approach to dealing with this proteome coverage problem is to prefractionate samples, either as proteins prior to enzymatic digestion (Alternate Protocol 3, Basic Protocol 4) or as peptides prior to MS analysis (Basic Protocol 7). To further increase the depth of LCMS/MS analysis of already fractionated samples, the authors have found that re-analyzing each sample, but excluding previously analyzed components, can increase the number of proteins identified by up to 20%. Using the exclude function, common on most new MS instruments, one can access components in the sample that the instrument did not have time to analyze during the first pass through. The other concern that arises when intent on identifying proteins of “biological interest” is the limit of detection of the MS instrumentation. By factoring in the limit of detection, the total amount of protein needed for a successful identification can be estimated. For proteins in a complex mixture, assume a general MS limit of detection to be at least 1 picomole (10–12 moles). For example, isolation and identification of a 50-kDa phosphoprotein from a cell lysate would require at least 50 ng of that protein present to maximize the chance of detecting it. Consequently, using 50 µg of cell lysate to identify this protein would assume that the phosphoprotein of interest makes up 0.1% of the entire lysate (which is not likely). Starting with 0.5 to 1 mg of cell lysate would maximize the chance of successfully identifying that protein. Much effort has been put into the development of proteomic methods using model organisms and standardized cell culture systems. Currently, the state of hESC derivation, culture, and differentiation is far from standardized. As such, using the hESC system to

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develop new proteomic approaches is not recommended. The authors suggest addressing a particular experimental question based on proteomics methods already established in model systems, adapting them to hESCs using the generic proteomic protocols provided in this unit. This will not only give the best procedural guideline, but also provide a benchmark with which to compare the results.

Anticipated Results The best indicator for the success of a proteomic analysis is in the number and/or types of proteins that are identified. Because there are different experimental strategies, these results can vary widely. In the case of hESCs, success should be measured by comparing the results of the proteomic study to the original methodologies after which the work is modeled. For example, imagine undertaking a study to identify membrane proteins. On standard tissue culture cells the standard methodology identifies ∼500 unique proteins, but when the exact same analysis is performed using hESCs, only ∼50 are identified. This suggests one or more problems have occurred: (1) not enough protein was used, (2) a correct/thorough enough MS analysis was not performed, or (3) the data interpretation methods may be flawed (e.g., incorrect parameters used for the database search). While identification of a single protein can be accomplished quickly and economically by mass spectrometry, larger-scale, high-throughput experiments require a lot of starting material (milligrams of protein for PTM analysis) and up to hundreds of hours of MS instrument time. For that reason, using more sample, more thorough sample fractionation, and more instrument and analysis time will ultimately lead to experimental results of higher quality.

Time Considerations Time requirements for proteomic experiments can vary depending on the type of experiment, level of analysis, or number of proteins to be identified. For details on time requirements for specific procedures, optional stopping points, and storage of samples during stopping points, refer to the instructions provided in the individual protocols within this unit.

Proteomic Analysis of Pluripotent Stem Cells

Preparation of hESCs Typical experiments can require anywhere from 106 to 108 hESCs depending on whether the goal is to identify a single protein or a multitude of post-translational modifications.

As such, starting with a single 9.8-cm2 well of a 6-well plate of ∼2 × 106 hESC and passaging 1:2 every 5 to 8 days, it could take between 1 and 6 weeks to obtain the number of cells necessary to start a particular experiment. For more details on hESC culture time requirements and expected cell number see the other hESC units in this manual. Protein extraction and evaluation Extraction of protein from hESCs (Basic Protocol 1, Alternate Protocol 1, 2, or 3) and subsequent evaluation of the extraction by protein assay (Basic Protocol 3) can be easily accomplished in a single day, taking into account preparation of extraction buffers that day. However, some of the various protease and phosphatase inhibitors, as well as nucleases can take 2 to 3 hr to prepare by themselves and should be made up ahead of the protein extraction. Moreover, affinity based applications such as purification of protein complexes or proteins based on PTMs can take additional time. In the authors’ experience, many immunoprecipitation procedures are carried out overnight and can add at least another day to a procedure. Gel electrophoresis A standard SDS-PAGE can be prepared and run in as little as 3 to 4 hr if the samples are already ready dissolved in 1× Laemmli buffer. For 2-D gel electrophoresis, the IEF step will take 2.5 to 3 days where the IPG strip is rehydrated with protein extract the first night and focused the second night. The second dimension SDS-PAGE, depending on the size, can take another 0.5 to 1 day to make and run (Basic Protocol 4). Staining of 1D or 2-D gels by colloidal Coomassie staining (Support Protocol 2) can take up to another 3 days, while standard Coomassie or Sypro staining still takes 2 days for completion. Digestion with trypsin Digestion of proteins from PAGE-based samples (Basic Protocol 5) generally takes 3 days to complete. Gel pieces are divided and cut up on the first day, stained, and then destained overnight. On the second day, samples are reduced and alkylated and then digested overnight with trypsin. On the third day, peptides are extracted and then dried for analysis by MS. Proteins in solution can be digested (Alternate Protocol 5) and ready for MS analysis in 2 days. In this case, samples are reduced and alkylated on the first day and allowed to

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digest overnight with trypsin. The following day, tryptic peptides are extracted and desalted (Basic Protocol 6), then dried down prior to MS analysis. If protein samples in solution are to be concentrated or bufferexchanged prior to digestion (Support Protocol 1), an extra day will be added to the procedure. Analysis by mass spectrometry Analysis time by mass spectrometry can be highly variable depending on the nature of the sample and instrumentation. Once digested, identifying a single protein by MALDI-TOF MS can be completed in 1 hr, including analysis, data processing, and database identification. However, for more complex samples where LC-MS/MS identification is necessary, the instrument analysis by itself can take an hour or more per sample, not including instrument calibration and conditioning. Moreover, this type of data takes a lot longer to process post-analysis and is much more complicated to interpret. Consequently, a basic analysis of a complex digest fractionated into 12 parts (Basic Protocol 7) prior to analysis by LC-MS/MS can take up to a week, including instrument, processing, and interpretation time.

Literature Cited Adachi, J., Kumar, C., Zhang, Y., Olsen, J.V., and Mann, M. 2006. The human urinary proteome contains more than 1500 proteins, including a large proportion of membrane proteins. Genome Biol. 7:R80. Aebersold, R. and Mann, M. 2003. Mass spectrometry-based proteomics. Nature 422:198-207. Andersen, J.S. and Mann, M. 2006. Organellar proteomics: Turning inventories into insights. EMBO Rep. 7:874-879. Andersen, J.S., Lam, Y.W., Leung, A.K., Ong, S.E., Lyon, C.E., Lamond, A.I., and Mann, M. 2005. Nucleolar proteome dynamics. Nature 433:7783. Beck, H.C., Nielsen, E.C., Matthiesen, R., Jensen, L.H., Sehested, M., Finn, P., Grauslund, M., Hansen, A.M., and Jensen, O.N. 2006. Quantitative proteomic analysis of post-translational modifications of human histones. Mol. Cell Proteomics 5:1314-1325. Blagoev, B. and Mann, M. 2006. Quantitative proteomics to study mitogen-activated protein kinases. Methods 40:243-250. Blagoev, B., Ong, S.E., Kratchmarova, I., and Mann, M. 2004. Temporal analysis of phosphotyrosine-dependent signaling networks by quantitative proteomics. Nat. Biotechnol. 22:1139-1145. Candiano, G, Bruschi, M., Musante, L., Santucci, L., Ghiggeri, G.M., Carnemolla, B., Orecchia, P., Zardi, L., and Righetti, P.G. 2004. Blue silver: A very sensitive colloidal Coomassie G-250

staining for proteome analysis. Electrophoresis 25:1327-1333. Chen, E.I., Hewel, J., Felding-Habermann, B., and Yates, J.R. 3rd. 2006. Large scale protein profiling by combination of protein fractionation and multidimensional protein identification technology (MudPIT). Mol. Cell Proteomics 5:5356. Delahunty, C. and Yates, J.R. 3rd. 2005. Protein identification using 2-D-LC-MS/MS. Methods 35:248-255. Domon, B. and Aebersold, R. 2006. Mass spectrometry and protein analysis. Science 312:212-217. Foster, L.J., de Hoog, C.L., Zhang, Y., Xie, X., Mootha, V.K., and Mann, M. 2006. A mammalian organelle map by protein correlation profiling. Cell 125:187-199. Gallagher, S.R. 2006. One-Dimensional SDS Gel Electrophoresis of Proteins. Curr. Protoc. Mol. Biol. 75:10.2A.1-10.2A.37. Gavin, A.C., Bosche, M., Krause, R., Grandi, P., Marzioch, M., Bauer, A., Schultz, J., Rick, J.M., Michon, A.M., Cruciat, C.M., Remor, M., Hofert, C., Schelder, M., Brajenovic, M., Ruffner, H., Merino, A., Klein, K., Hudak, M., Dickson, D., Rudi, T., Gnau, V., Bauch, A., Bastuck, S., Huhse, B., Leutwein, C., Heurtier, M.A., Copley, R.R., Edelmann, A., Querfurth, E., Rybin, V., Drewes, G., Raida, M., Bouwmeester, T., Bork, P., Seraphin, B., Kuster, B., Neubauer, G., and Superti-Furga, G. 2002. Functional organization of the yeast proteome by systematic analysis of protein complexes. Nature 415:141-147. Glish, G.L. and Vachet, R.W. 2003. The basics of mass spectrometry in the twenty-first century. Nat. Rev. Drug Discov. 2:140-150. Golemis, E.A., Serebriiskii, I., Finley, R.L.Jr., Kolonin, M.G., Jeno Gyuris, J., and Brent, R. 1998. Interaction trap/two-hybrid system to identify interacting proteins. Curr. Protoc. Prot. Sci. 14:19.2.1-19.2.40. Ho, Y., Gruhler, A., Heilbut, A., Bader, G.D., Moore, L., Adams, S.L., Millar, A., Taylor, P., Bennett, K., Boutilier, K., Yang, L., Wolting, C., Donaldson, I., Schandorff, S., Shewnarane, J., Vo, M., Taggart, J., Goudreault, M., Muskat, B., Alfarano, C., Dewar, D., Lin, Z., Michalickova, K., Willems, A.R., Sassi, H., Nielsen, P.A., Rasmussen, K.J., Andersen, J.R., Johansen, L.E., Hansen, L.H., Jespersen, H., Podtelejnikov, A., Nielsen, E., Crawford, J., Poulsen, V., Sorensen, B.D., Matthiesen, J., Hendrickson, R.C., Gleeson, F., Pawson, T., Moran, M.F., Durocher, D., Mann, M., Hogue, C.W., Figeys, D., and Tyers, M. 2002. Systematic identification of protein complexes in Saccharomyces cerevisiae by mass spectrometry. Nature 415:180-183. Hoffman, L.M. and Carpenter, M.K. 2005. Characterization and culture of human embryonic stem cells. Nat. Biotechnol. 23:699-708. Kirkpatrick, D.S., Denison, C., and Gygi, S.P. 2005a. Weighing in on ubiquitin: The expanding role of mass-spectrometry-based proteomics. Nat. Cell Biol. 7:750-757.

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Kirkpatrick., D.S., Gerber, S.A., and Gygi, S.P. 2005b. The absolute quantification strategy: A general procedure for the quantification of proteins and post-translational modifications. Methods 35:265-273.

Ranish, J.A., Yi, E.C., Leslie, D.M., Purvine, S.O., Goodlett, D.R., Eng, J., and Aebersold, R. 2003. The study of macromolecular complexes by quantitative proteomics. Nat. Genet. 33:349355.

Kislinger, T., Cox, B., Kannan, A., Chung, C., Hu, P., Ignatchenko, A., Scott, M.S., Gramolini, A.O., Morris, Q., Hallett, M.T., Rossant, J., Hughes, T.R., Frey, B., and Emili, A. 2006. Global survey of organ and organelle protein expression in mouse: Combined proteomic and transcriptomic profiling. Cell 125:173186.

Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular cloning: A laboratory manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

Kratchmarova, I., Blagoev, B., Haack-Sorensen, M., Kassem, M., and Mann, M. 2005. Mechanism of divergent growth factor effects in mesenchymal stem cell differentiation. Science 308:1472-1477.

Steen, H. and Mann, M. 2004. The ABC’s (and XYZ’s) of peptide sequencing. Nat. Rev. Mol. Cell Biol. 5:699-711.

Krokhin, O.V., Craig, R., Spicer, V., Ens, W., Standing, K.G., Beavis, R.C., and Wilkins, J.A. 2004. An improved model for prediction of retention times of tryptic peptides in ion pair reversedphase HPLC: Its application to protein peptide mapping by off-line HPLC-MALDI MS. Mol. Cell Proteomics 3:908-919. Mann, M. 2006. Functional and quantitative proteomics using SILAC. Nat. Rev. Mol. Cell Biol. 7:952-958. Mann, M. and Jensen, O.N. 2003. Proteomic analysis of post-translational modifications. Nat. Biotechnol 21:255-261. McCarthy, J., Hopwood, F., Oxley, D., Laver, M., Castagna, A., Righetti, P.G., Williams, K., and Herbert, B. 2003. Carbamylation of proteins in 2-D electrophoresis–myth or reality? J. Proteome Res. 2:239-242. Ong, S.E. and Mann, M. 2005. Mass spectrometrybased proteomics turns quantitative. Nat. Chem. Biol. 1:252-262. Ong, S.E., Blagoev, B., Kratchmarova, I., Kristensen, D.B., Steen, H., Pandey, A., and Mann, M. 2002. Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol. Cell Proteomics 1:376-386. Pedrioli, P.G., Eng, J.K., Hubley, R., Vogelzang, M., Deutsch, E.W., Raught, B., Pratt, B., Nilsson, E., Angeletti, R.H., Apweiler, R., Cheung, K., Costello, C.E., Hermjakob, H., Huang, S., Julian, R.K., Kapp, E., McComb, M.E., Oliver, S.G., Omenn, G., Paton, N.W., Simpson, R., Smith, R., Taylor, C.F., Zhu, W., and Aebersold, R. 2004. A common open representation of mass spectrometry data and its application to proteomics research. Nat. Biotechnol. 22:14591466. Ptacek, J. and Snyder, M. 2006. Charging it up: Global analysis of protein phosphorylation. Trends Genet. 22:545-554.

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Raman, R., Raguram, S., Venkataraman, G., Paulson, J.C., and Sasisekharan, R. 2005. Glycomics: An integrated systems approach to structure-function relationships of glycans. Nat. Methods 2:817-824.

Schmelzle, K. and White, F.M. 2006. Phosphoproteomic approaches to elucidate cellular signaling networks. Curr. Opin. Biotechnol. 17:406414.

Stelzl, U., Worm, U., Lalowski, M., Haenig, C., Brembeck, F.H., Goehler, H., Stroedicke, M., Zenkner, M., Schoenherr, A., Koeppen, S., Timm, J., Mintzlaff, S., Abraham, C., Bock, N., Kietzmann, S., Goedde, A., Toksoz, E., Droege, A., Krobitsch, S., Korn, B., Birchmeier, W., Lehrach, H., and Wanker, E.E. 2005. A human protein-protein interaction network: A resource for annotating the proteome. Cell 122:957-968. Wang, L., Li, L., Menendez, P., Cerdan, C., and Bhatia, M. 2005. Human embryonic stem cells maintained in the absence of mouse embryonic fibroblasts or conditioned media are capable of hematopoietic development. Blood 105:45984603. Wang, J., Rao, S., Chu, J., Shen, X., Levasseur, D.N., Theunissen, T.W., and Orkin, S.H. 2006. A protein interaction network for pluripotency of embryonic stem cells. Nature 444:364-368. Washburn, M.P., Wolters, D., and Yates, J.R. 3rd. 2001. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19:242-247. Wu, C.C. and Yates, J.R. 3rd. 2003. The application of mass spectrometry to membrane proteomics. Nat. Biotechnol. 21:262-267. Wu, C.C., MacCoss, M.J., Howell, K.E., and Yates, J.R. 3rd. 2003. A method for the comprehensive proteomic analysis of membrane proteins. Nat. Biotechnol. 21:532-538. Xu, C., Inokuma, M.S., Denham, J., Golds, K., Kundu, P., Gold, J.D., and Carpenter, M.K. 2001. Feeder-free growth of undifferentiated human embryonic stem cells. Nat. Biotechnol. 19:971-974.

Key References Aebersold, R. and Mann, M. 2003. See above. Domon, B. and Aebersold, R. 2006. See above. Steen, H. and Mann, M. 2004. See above. General overviews of proteomics. They contain brief descriptions of current applications, popular types of instrumentation, and information on peptide sequencing and identification. Glish, G.L. and Vachet, R.W. 2003. See above. Basic review of mass spectrometry theory, instrumentation, and application.

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Internet Resources http://www.bio-rad.com/LifeScience/pdf/ Bulletin 9004.pdf Bio-Rad Protein assay application manual and compatibility information http://www1.amershambiosciences.com/aptrix/ upp01077.nsf/Content/2-D electrophoresis∼2Delectrophoresis handbook GE Healthcare two-dimensional gel electrophoresis application and troubleshooting guide. http://www.bioinfor.com:8080/peaksonline http://www.thegpm.org http://www.matrixscience.com/ search form select.html Free online search tools for proteomic data. http://www.proteomecommons.org/ http://www.expasy.org/tools/ http://tools.proteomecenter.org/software.php Repository of proteomic software and information. http://www.biomart.org/ Tool for parsing and annotating large protein datasets (i.e., convert protein accession numbers to gene accession number, or add gene ontology information to a dataset). http://www.ionsource.com/ http://www.spectroscopynow.com/ General proteomic and mass spectrometry information.

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Gene Expression Analysis of RNA Purified from Embryonic Stem Cells and Embryoid Body–Derived Cells Using a High-Throughput Microarray Platform

UNIT 1B.2

Audrey Player,1 Yonghong Wang,2 Mahendra Rao,3 and Ernest Kawasaki1 1

National Cancer Institute, Bethesda, Maryland SAIC-Frederick, Inc., NCI-Frederick, Frederick, Maryland 3 Johns Hopkins University School of Medicine, Baltimore, Maryland and Invitrogen Corp. Carlsbad California, 2

ABSTRACT In this unit, starting with purified RNA, experimental protocols for performing microarrray expression analysis of embryonic stem cell lines compared to their corresponding differentiated embryoidal bodies are described. Methods for data analysis are suggested, with the goal of determining which genes are differentially expressed between the preparations. As an example, the use of the Affymetrix microarray expression platform is described, but alternative experimental options for analysis of RNA transcript levels are also summarized. This unit suggests quality control metrics, summazrizes the critical parameters necessary for obtaining reproducible experimental results, and outlines quantitative PCR methods for validating microarray results. Curr. Protoc. Stem Cell Biol. C 2007 by John Wiley & Sons, Inc. 2:1B.2.1-1B.2.36.
Keywords: embryonic stem cells r embryoid bodies r RNA expression analysis r microarray platform

INTRODUCTION Embryonic stem (ES) cell types are derived from pluripotent cells isolated from the inner cell mass of pre-implantation embryos. ES cells will differentiate if allowed to aggregate (Doetschman et al., 1985) or if incubated in differentiation media (Ohlsson et al., 1993; Shamblott et al., 1998; Itskovitz-Eldor et al., 2000) leading to the formation of embryoid bodies (EB). Understanding the genetic changes that occur in the reprogramming from ES to EB cells has been useful in helping to characterize stem cells. Genes and pathways important in the programming of ES cells to the differentiated EB cell types have been identified by a number of investigators (Grabel et al., 1998; Miura et al., 2004; Bhattacharya et al., 2005; Skottman et al., 2005; Wei et al., 2005), many using global RNA expression microarray platforms. Unlike its predecessors, northern blots or PCR-based platforms, RNA expression microarrays allow for the simultaneous interrogation of thousands of transcripts per a single biological sample, which is critical for a more complete understanding of a very complex process. Many of the current commercially available microarray platforms contain the full complement of the human genome, as characterized by the Human Genome sequencing efforts (Lander et al., 2001); this includes the non-redundant, known genes or reference sequences (RefSeq: ftp://ftp.ncbi.nih.gov/refseq/), and the predicted-novel genes or expressed sequence tags (ESTs: http://hgdownload.cse.ucsc.edu/goldenPath/hg18/bigZips/). This unit details gene expression analysis in embryonic stem cell (ESC) RNA samples using the Affymetrix microarray platform. As an example, data generated by comparing

Current Protocols in Stem Cell Biology 1B.2.1-1B.2.36 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b02s2 C 2007 John Wiley & Sons, Inc. Copyright


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two embryonic stem cell samples to their corresponding embryoid body–derived samples are used. Culture conditions for ESC and EB have been described previously (Bhattacharya et al., 2004, 2005). RNA is purified using Trizol (http://www. Invitrogen.com; Bhattacharya et al., 2004; Liu et al., 2006), but alternative methods include RNeasy (http://www.Qiagen.com) or RNA Stat (http://www.Tel-Test.com), just to name a few. There are a vast number of commercially available kits, that depend on the source (i.e., tissue culture, blood, etc.) and quantity of the biological sample. The quality of the samples used in these protocls has been previously validated by examining the presence of early markers of differentiation (Cai et al., 2006; Liu et al., 2006; Loring and Rao, 2006; Shin and Rao, 2006; see Support Protocols 1 and 2). This is critical in determining which samples are of adequate quality for further analysis or for pooling of data in multivariate comparisons. If the investigator is not limited by the amount of RNA sample available, any platform can be used (Jarvinen et al., 2004; Ball et al., 2005; Li et al., 2005; de Reynies et al., 2006). The authors chose the Affymetrix Genechip microarray platform (which is a short oligonucleotide platform) for assessment of the RNA expression levels of the samples. The Affymetrix Genechip microarray is a single-color array, therefore, for one sample, one Genechip is used. The platform requires that total RNA be amplified, and hybridized to the Genechip as biotin-modified antisense RNA (aRNA). Starting with purified ESC and EB RNA, protocols used in the authors’ laboratory are described for (1) RNA amplification including generation of biotin-modified aRNA (see Basic Protocol 1 and Alternate Protocol 1); (2) hybridization of the stem cell RNA to the Genechip microarray; (3) wash, stain, and detection of signal intensities (see Basic Protocol 3), which allow for determination of relative transcript expression levels; (4) preliminary statistical analysis of the microarray results (see Basic Protocol 4 and Alternate Protocol 2), and (5) validation of the results using real-time PCR (see Support Protocol 3). Regardless of the microarray platform used, the reproducibility, hence reliability of the results, critically depends on establishing stringent quality metrics for various steps in the protocol. This requires that the entire protocol be standardized. For example, parameters designating the quality and concentration of the RNA starting material, and its amplification must be established and consistent for the duration of the study. All hybridizations must be performed under as similar as possible conditions with technical and biological replicates, and only microarrays with comparable quality metrics analyzed and compared.

STRATEGIC PLANNING One should first consider the quantity of total RNA input to use for the study to ensure equivalent mRNA representation per sample. As part of the microarray protocol, RNA is amplified to generate adequate amounts for application to the Genechip microarray and for incorporation of modified nucleotides for detection of transcripts. The total RNA input concentration also determines whether one or two cycles of RNA amplification is performed. One complete amplification cycle starting with total RNA input results in the generation of first strand cDNA, second strand cDNA, and aRNA. Two-cycle amplification entails one complete amplification cycle followed by the use of the amplication product as template for another complete cycle. Higher concentrations of RNA input require one cycle, and lower concentrations require two cycles of amplification; details of the requirements are specified in Critical Parameters. For the current study, 1 µg of total RNA input isolated from 106 cultured cells is used and one cycle amplification is performed. Microarray Analysis of Stem Cell Gene Expression

A commercial source for the RNA amplification kit must also be considered. Once a particular amplification kit is chosen, it should be used for the duration of the study, as different kits can produce somewhat different products, and inconsistent experimental results. There are a number of commercially available RNA amplification kits that can be used to

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generate modified aRNA necessary for hybridization to the Affymetrix Genechip. Ambion (http://www.ambion.com; http://www.appliedBiosystems.com), Enzo (http://www. enzo.com), Epicentre (http://www.epibio.com), Genisphere (http://www.genisphere.com), Invitrogen (http://www.invitrogen.com), Kreatech Biotechnology (http://www.kreatech. com), Roche (http://www.roche.com), and SuperArray (http://www.superarray.com) are a few of the companies that manufacture RNA amplification kits. The reagents indicated are for research purposes only. When choosing an aRNA amplification kit, make sure the product is compatible for hybridization to the platform of interest. For the current study, the Ambion MessageAmp II biotin-enhanced aRNA amplification kit (Ambion cat. no. 1791) is used with one cycle amplification, but the protocol also outlines twocycle amplification combined with use of the MessageAmp II aRNA kit (Ambion cat. no. 1751).

GENERATING cDNA AND IN VITRO–TRANSCRIPTION (IVT) aRNA For this protocol, 1 µg of ESC or EB total RNA is used and one-cycle aRNA amplification using the Ambion MessageAmp II biotin-enhanced kit is performed; however, the procedures for performing both one- and two-cycle aRNA amplification are outlined below. These instructions are adapted from the Ambion MessageAmp II kit protocols.

BASIC PROTOCOL 1

Materials ESC or EB RNA, quality assessed (see Support Protocol 1) Ambion MessageAmp II biotin-enhanced kit (cat. no. 1791) containing: T7 oligo(dT) primer Arrayscript reverse transcription enzyme RNAase inhibitor 10× first-strand buffer dNTP mixture 10× second-strand buffer DNA polymerase RNAase H T7 enzyme mixture T7 10× reaction buffer Biotin-UTP mixture (which also includes unmodified NTPs) Distilled water Ambion MessageAmp II kit components (Optional; cat. no. 1751). Same as above except different in vitro transcription reagents, and random hexamer (day 2 reagent) as noted below: T7 enzyme mixture T7 10× reaction buffer 75 mM T7 ATP solution 75 mM T7 CTP solution 75 mM T7 GTP solution 75 mM T7 UTP solution Random hexamer primer Distilled water Wash buffer (100% ethanol must be added before use) cDNA binding buffer aRNA binding buffer aRNA filter cartridges aRNA 1.7-ml collection tubes cDNA filter cartridges cDNA 1.7-ml collection tubes 5× fragmentation buffer 100% ethanol, ACS-grade

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1 M Tris-acetate, pH 8.1 (Trizma base, Sigma cat. no. T1503, pH adjusted with glacial acetic acid, Sigma cat. no. A628) Magnesium acetate (Sigma cat. no. M2545) Potassium acetate (Sigma cat. no. P5708) DEPC-treated water Nuclease-free 0.5-ml tubes, non-stick, sterile Vortex Microcentrifuge Thermal cycler (with bonnet), incubator, or temperature block with adjustable temperatures Vacuum centrifuge Vacuum dryer Spectrophotometer Agilent Bioanalyzer or gel electrophoresis equipment, optional 0.2-µm vacuum filter unit Synthesize first-strand cDNA 1. Place up to 10 µl of 1 µg total ESC or EB RNA into a sterile, nuclease-free 0.5-ml tube. RNA must be in high-quality water or low TE buffer. Up to 2 µg can be used with this amplification kit.

2. Add 1 µl of T7 oligo(dT) primer. 3. Add nuclease-free water to a final volume of 12 µl, vortex, mix, and then centrifuge to collect content at bottom of tube. 4. Incubate 10 min at 70◦ C. 5. Centrifuge samples briefly (∼5 sec) to collect content. Place mixture on ice. 6. At room temperature, prepare reverse transcription master mix in a nuclease-free tube. Assemble enough to synthesize first-strand cDNA for all RNA samples in the experiment including an excess of 5% to account for pipetting errors. The following amounts are per individual sample. Assemble the reverse transcription mixture in the order shown:

2 µl 10× first strand buffer 4 µl dNTP mixture 1 µl RNAse inhibitor 1 µl ArrayScript enzyme. 7. Mix well by gently vortexing or by finger flicking the tube. Centrifuge briefly (5 sec) to collect the reverse transcription master mix at the bottom of the tube and place at room temperature. 8. Transfer 8 µl of the reverse transcription mix to each total RNA sample. Mix thoroughly by pipetting up and down two to three times; centrifuge briefly to collect the sample. Total volume in tube should be 20 µl.

Microarray Analysis of Stem Cell Gene Expression

9. Place samples 2 hr in a 42◦ C incubator (use an incubator where condensation does not accumulate).

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Synthesize second-strand cDNA 10. Before completion of the first-strand reaction, prepare the second-strand master mix in a nuclease-free tube. Prepare enough for all RNA samples. Volumes below are per individual sample: 63 µl nuclease-free water 10 µl 10× second-strand buffer 4 µl dNTP mixture 2 µl DNA polymerase 1 µl RNase H. Mix gently. Centrifuge briefly to collect. Place on ice until ready to begin secondstrand synthesis. 11. Transfer 80 µl of second-strand master mix to each sample. Mix thoroughly by pipetting up and down. Centrifuge to collect content at bottom of tube. Total volume of this mixture should be 100 µl.

12. Place the tubes on a 16◦ C refrigerated block. Make sure the block is 16◦ C prior to placing samples. Incubate 2 hr. 13. Place samples on ice and proceed to cDNA purification.

Purify cDNA 14. Assemble cDNA purification cartiridge in a kit-provided wash tube. 15. Add 250 µl of the cDNA binding buffer to each sample. Mix well and apply to center of purification column. 16. Centrifuge 1 min at 10,000 × g (∼10,000 rpm), room temperature, in a microcentrifuge to bind cDNA to column. 17. Discard the flow-through and replace the cDNA filter cartridge in the wash tube (wash all cartridges using the same wash tube). 18. Apply 500 µl wash buffer to each cDNA filter cartridge. The wash solution comes as a 6-ml solution from the manufacturer; before using, add 24 ml of 100% ethanol.

19. Centrifuge 1 min at 10,000 × g (∼10,000 rpm), room temperature, to wash. 20. Discard the flow-through and spin the cDNA filter cartridge for an additional 1 min to remove trace amounts of wash buffer. 21. Transfer cDNA filter cartridge to a clean cDNA collection tube. 22. Apply 12 µl of prewarmed to 50◦ to 55◦ C nuclease-free water or low TE buffer to the center of the cDNA filter cartridge. Samples can be eluted in prewarmed (50◦ to 55◦ C) nuclease-free water or low TE buffer. If the samples are to be stored for times beyond a couple of days, the recommendation is to elute in low TE buffer.

23. Leave ∼2 min at room temperature, and then centrifuge 1.5 min at 10,000 × g (∼10,000 rpm), room temperature. Keep the eluant. 24. Repeat elution step 23 by adding 12 µl of nuclease-free water to the center of the filter cartridge and centrifuging. Keep the eluant. Sample should be in a final volume of ∼20 µl. If more, then concentrate using the vacuum dryer.

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This is a potential stopping point. Sample can be stored overnight at −20◦ C, or proceed directly to in vitro transcription step.

In vitro transcribe (IVT) samples using modified NTPs 25. At room temperature, prepare the IVT master mix by adding the following reagents to a nuclease-free tube in the order listed below. Volumes below are per individual sample: 12 µl biotin-NTP mixture (including unmodified NTPs and biotin-UTP) 4 µl T7 10× reaction buffer 4 µl T7 enzyme mix. 26. Mix well and collect. Add 20 µl of IVT master mix to each 20-µl cDNA sample. 27. Incubate 4 hr at 37◦ C. Proceed to aRNA purification.

Purify aRNA 28. Preheat nuclease-free elution water or low TE buffer to 50◦ to 55◦ C. Leave on heat block until ready. 29. Assemble aRNA filter cartridge in the wash tube. 30. Bring ESC and EB sample volume (from step 27) up to 100 µl with nuclease-free water, if necessary. 31. Add 350 µl of aRNA binding buffer to each sample. 32. Mix well by tapping tube with finger. 33. Add 250 µl of ACS-grade 100% ethanol to each sample. Mix well by tapping tube with finger. Proceed directly to the next step. CAUTION: Do not vortex and do not centrifuge.

34. Pipet the sample mixture onto the center of the filter cartridge. 35. Centrifuge for 1 min at 10,000 × g (∼10,000 rpm), room temperature. 36. Discard the flow-through and replace the filter cartridge in the same tube (wash all cartridges using the same wash tube). 37. Add 650 µl of the wash buffer to each cartridge. 38. Centrifuge 1 min at 10,000 × g (∼10,000 rpm), room temperature. Discard flowthrough and centrifuge for an additional 1 min to remove trace amounts of wash solution. 39. Place cartridge in a clean nuclease-free 1.7-ml collection tube. 40. Add up to 100 µl of nuclease-free water or low TE buffer (preheated to 50◦ to 55◦ C) onto the center of the cartridge. If the aRNA is to be stored for a couple of days, store aRNA as a concentrated sample in low TE buffer. This is done by eluting with low TE buffer then concentrating the sample using vacuum centrifugation (i.e., drying the sample in a vacuum dryer).

41. Leave for 2 min.

Microarray Analysis of Stem Cell Gene Expression

42. Centrifuge 1.5 hr at 10,000 × g (∼10,000 rpm), room temperature, to collect aRNA sample. Keep the eluant. 43. Repeat elution step by adding up to 100 µl of nuclease-free water (preheated to 50◦ to 55◦ C) onto the center of the cartridge.

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44. Leave on cartridge for 2 min. 45. Centrifuge for 1.5 min at 10,000 × g (∼10,000 rpm), room temperature, to collect aRNA sample. Keep the eluant (final volume ∼200 µl). Determine the RNA concentration using a spectrophotometer (see Assessing RNA yield below.) Alternatively, examine RNA profile and determine concentration using the Bioanalyzer (see Support Protocol 1). It is important to determine the aRNA concentration at this point because the “beforefragmentation-aRNA concentration” is used for calculating the amount hybridized to the Genechip. An example of the aRNA Bioanalyzer profile is given in Figure 1B.2.1, lane 3. One cycle aRNA should be ∼1 kb in length.

46. aRNA can be concentrated, if needed and fragmented (see below) for application to the Genechip microarray. If a one-cycle reaction is used, then proceed to step 48 below.

Determine aRNA yield by absorbance 47. Calculate the aRNA yield. For example: aRNA is in 100 µl 2 µl of the prep is diluted 1:10 into 18 µl of low TE buffer or nuclease-free water A260 = 0.75 RNA concentration = 0.75 × 10 × 40 µg/ml = 300 µg/ml or 0.3 µg/µl 98 µl of the aRNA remaining: 98 µl × 0.3 µg/µl = 29.4 µg. Fragment aRNA Genes on the Affymetrix platform are represented by ten to twenty 25-mer oligonucleotides (or probes) tiled to represent the complimentary sequence of a particular gene or target. Collectively, these probes are referred to as probe-sets. A probe-set corresponds to one gene or target. Specific oligonucleotides or probes within a feature on the Affymetrix microarray Genechips are short 25-mers, separated by distances measured ◦ in Angstroms. To allow for efficient hybridization and elimination of steric constraints, aRNA is fragmented by heating 35 min at 95◦ C in fragmentation buffer. 48. To prepare 5× fragmentation buffer, combine the following components to a total volume of 20 ml:

4.0 ml 1 M Tris-acetate, pH 8.1 0.64 g magnesium acetate 0.98 g potassium acetate DEPC-treated water to 20 ml. Mix thoroughly and filter through a 0.2-µm vacuum filter unit. Aliquot and store at room temperature. The 5× fragmentation buffer is frequently depleted. Prepare an ample amount.

49. Measure the absorbance of the aRNA. Sample concentrations should be at least 0.2 µg/µl; if lower, concentrate using a vacuum dryer.

50. Prepare the following mixture:

aRNA (15 to 20 µg) in up to 32 µl 8 µl 5× fragmentation buffer Nuclease-free water to 40 µl.

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Figure 1B.2.1 RNA profile analysis. Examples of the RNA profiles are demonstrated. Lane 1: RNA ladder used as a size marker. Lane 2: Total RNA profile demonstrating size of the 28S (∼5 kilobases (kb) and 18S (∼2 kb) ribosomal bands. Lane 3: aRNA profile following one cycle of amplification. Lane 4: aRNA profile following two cycles of amplification. Lane 5: aRNA following fragmentation.

51. Incubate 35 min at 95◦ C. Centrifuge to collect content at bottom of tube, and then place on ice. 52. Remove 2-µl aliquot for analysis. Fragmented aRNA size should range between 50 and 200 nucleotides. An example of the fragmented aRNA profile is given in Figure 1B.2.1, lane 5. Fragmented ESC or EB RNA is stable for up to 1 year at −80◦ C. SUPPORT PROTOCOL 1

ASSESSMENT OF ESC AND EB RNA QUALITY USING THE AGILENT BIOANALYZER The SA02 ESC (Heins et al., 2004) and differentiated EB, and the BG01 ESC (Mitalipova et al., 2003) and the differentiated EB are used to demonstrate microarray analysis in this section. ESC cells are cultured and EB cells generated and validated as described in previous studies (Bhattacharya et al., 2004, 2005; Cai et al., 2006; Liu et al., 2006; Loring and Rao, 2006; Shin and Rao, 2006). Before starting the experiment, it is important to examine the quality of the ESC and EB total RNA samples because poor-quality material will lead to inefficient amplification, and inconsistent results. The methods used to establish the quality metrics and the range of acceptable metrics are listed below. Determine the absorbance, A260/280 ratio of the samples using a spectrophometer or similar equipment. Quality RNA, free of DNA and protein contaminants, ranges from 1.8 to 2.1. Examine the RNA profile by separation on an agarose gel or Agilent Bioanalyzer. The ribosomal 28S (∼5 kb in size) and 18S (∼2 kb in size) bands should be prominent with ratios ranging from 2.0 to 1.4 (28S/18S; http://www.ambion.com/ techlib/basics/rnasecontrol/index.html). This is determined using the Agilent Bioanalyzer, which calculates the ratio automatically; when using agarose gels, this value is estimated.

Microarray Analysis of Stem Cell Gene Expression

The Bioanalyzer also calculates an RNA integrity number (RIN), which is an estimate of the integrity of the RNA sample. Samples with RIN numbers ranging from 10.0 to 7.0 are

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considered good-quality RNA. The RIN value is calculated based on the 28S/18S ratio and the amount of degraded species located at various positions along the RNA profile, which presents as additional bands along the length of the profile or lower-molecularweight species near the bottom of the profile. The authors use the Agilent Bioanalyzer, and instructions (modified from the manufacturer) on its use are outlined below.

Materials 1 µg RNA isolated from ∼106 ESC and EB cells Agilent Nanochip, Lab-on-a chip (cat. no. 5067-1511) including: Detection chips Cleaning chips Syringe Gel matrix RNA ladder Spin filters Gel dye concentrate NanoMarker Nuclease-free water Low TE buffer (10 mM Tris·Cl, pH 7.4 and 1 mM EDTA) RNaseZap (Ambion cat. no. 9780) Agilent Bioanalyzer 2100 consisting of: Priming station Bioanalyzer vortexer 70◦ C heating block Centrifuge 1.5-ml nuclease-free tubes Prepare ESC and EB samples and RNA ladder 1. Heat-denature 2 µl each of the ESC and EB total RNA samples and 2 µl of the RNA ladder for 2 min at 70◦ C. The sample should be in nuclease-free water or low TE buffer. A 1-µl volume of RNA sample will be used, but 2 µl is denatured to compensate for recovery issues.

2. Quick cool on ice ∼30 sec and centrifuge to collect content at bottom of tube. Place at room temperature until ready for application to the detection chip. Use within 1 hr.

Decontaminate Bioanalyzer electrodes 3. Place 350 µl of RNaseZap into one of the cleaning chip wells. All wells are connected, therefore, 350 µl should be enough to disperse and fill the entire chip.

4. Place the cleaning chip (with RNaseZap) into the Bioanalyzer. Close the cover, and leave for ∼1 min. 5. Remove the cleaning chip. 6. Add 350 µl of water to another cleaning chip. Fill as above. 7. Place in the Bioanalyzer, and leave for 1 min. 8. Open the cover to allow drying (∼30 sec). The instrument is now ready for use.

Prepare the gel-dye mixture 9. Assemble the spin filter. Place 400 µl of RNA gel matrix (red top) into spin filter. 10. Centrifuge 10 min at 1500 × g, room temperature. Use within 4 weeks.

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11. Mix 130 µl filtered RNA gel matrix (from above) with 2 µl RNA dye concentrate (blue top) in a clean 1.5-ml nuclease-free tube. This is the working “gel dye mix.”

12. Vortex well. Store up to 4 weeks at 4◦ C protected from light.

Load gel dye mix onto the detection chip 13. Place a new detection chip on chip priming station. 14. Add 9 µl of the gel dye mix to the blackened well marked “G”, being careful not to introduce bubbles. 15. Make sure the plunger is at the 1-ml position and then close the chip priming station. 16. Apply pressure by pressing the plunger until it is held by the syringe chip (e.g., until a click is heard). 17. Wait for exactly 30 sec, and then release the plunger using the chip release mechanism. 18. Wait 30 sec then pull back the plunger to the 1-ml position. 19. Open the chip priming station. 20. Pipet 9 µl of gel dye mix into each of the other two wells marked “G”.

Load RNA Nanomarker, ladder, and ESC and EB samples to the detection chip 21. Pipet 5 µl of the RNA 6000 NanoMarker (green top) into all wells marked 1 to 12 (i.e., sample wells) and the well with the ladder designation. 22. Pipet 1 µl of the RNA 6000 ladder into the well with the ladder designation. 23. Pipet 1 µl of ESC or EB RNA samples into each of the 12 sample wells (labeled 1 through 12). If all 12 wells are not being used, then add 1 µl of water to the unused wells.

24. Place the chip on the Bioanalyzer vortexer and vortex for 1 min at the recommended set-point of 2400. 25. Run the chip within 10 min of loading and vortexing. 26. Place the chip in the Bioanalyzer and close the cover.

Run assay 27. Open the Agilent 2100 BioSizing program. From the top menu, choose the appropriate program to Run. In this case, designate Assay/RNA/Eukaryotic Total RNA Nano. Click Start to begin. 28. View the data by clicking View or Graph from the top menu. Example of the RNA ladder and good quality total RNA profile are given in Figure 1B.2.1, lanes 1 and 2, respectively. ALTERNATE PROTOCOL 1

Microarray Analysis of Stem Cell Gene Expression

TWO-CYCLE aRNA AMPLIFICATION If the quantity of RNA is low, two-cycle aRNA amplification is used to prepare sufficient material for hybridization. First and second strand generation is the same as previously noted (Basic Protocol 1), however, some volumes differ and for in vitro translation, unmodified NTPs are used.

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Additional Materials (also see Basic Protocol 1) cDNA (see Basic Protocol 1, steps 1 to 24) Ambion MessageAmp II (cat. no. 1751) or Ambion MessageAmp II biotin-enhanced (cat. no. 1791) Random hexamer Prepare aRNA 1. Synthesize first-strand cDNA, second-strand cDNA, and purify as noted in Basic Protocol 1, steps 1 to 24. Vacuum dry cDNA to 16 µl. 2. Prepare IVT master mix with unmodified NTPs as follows (component volumes below are per individual sample):

4 µl 75 mM T7 ATP solution 4 µl 75 mM T7 CTP solution 4 µl 75 mM T7 GTP solution 4 µl 75 mM T7 UTP solution 4 µl T7 10× reaction buffer 4 µl T7 enzyme mix. 3. Mix well and collect master mix by centrifugation. 4. Add 24 µl of the master mix to each 16 µl cDNA sample. 5. Mix by pipetting up and down. Collect content to bottom of tube by centrifuging. 6. Incubate 14 hr in a 37◦ C oven (do not allow condensation). 7. Perform aRNA purification (see Basic Protocol 1, steps 28 to 45). Final volume of aRNA should be ∼200 µl.

8. Vacuum dry the sample to ∼50 to 100 µl, then examine profile and determine RNA concentration using the Bioanalyzer (see Support Protocol 1). If aRNA concentration is undetectable, the sample should be concentrated (i.e., dried) to an even smaller volume. The aRNA can be stored overnight at −20◦ C, or proceed to two-cycle amplification.

Synthesize first and second strand 9. Collect 1 µg of aRNA in up to 10 µl of high-quality nuclease-free water in a 2-ml sterile, nuclease-free tube. Be careful to use the same concentrations for ESC or EB samples. This will require concentrating the aRNA sample, from cycle one, using a vacuum dryer. Up to 2 µg aRNA can be used with the Ambion MessageAmp kits.

10. Add 2 µl of second-round primers (random hexamer). 11. Bring up to 12 µl with nuclease-free water. 12. Incubate 10 min at 70◦ C. 13. Remove, collect by centrifuging ∼5 sec, and place on ice. 14. Prepare second-round, first-strand master mix as follows (volumes below are per individual samples):

2 µl 10× first-strand buffer 4 µl dNTP mix

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1 µl RNase inhibitor 1 µl ArrayScript enzyme. 15. Mix well by pipetting up and down. Centrifuge briefly (∼5 sec). 16. Add 8 µl of the master mix and mix by pipetting up and down. 17. Place samples 2 hr at 42◦ C. 18. Add 1 µl of RNase H to each sample, mix gently, and incubate 30 min 37◦ C. 19. Add 5 µl of T7 oligo(dT) primer to each sample. Incubate 10 min at 70◦ C. 20. Place ∼1 min on ice to quick cool. Collect content at bottom of tube by centrifugation. 21. Prepare second-strand master mix as follows (add 74 µl to each sample):

58 µl nuclease-free water 10 µl 10× second-strand buffer 4 µl dNTP 2 µl DNA polymerase. 22. Mix well. Centrifuge briefly. 23. Place sample 2 hr on 16◦ C refrigerated block. Make sure block is cooled to 16◦ C before adding samples.

24. Purify cDNA (see Basic Protocol, steps 14 to 24). Final elution volume should be 20 µl. If a larger volume is obtained, cDNA sample can be concentrated using the vacuum dryer.

In vitro transcribe (IVT) using modified NTPs 25. Perform IVT (see Basic Protocol 1, steps 25 to 27, except in step 27, incubate 14 hr at 37◦ C). Purify aRNA 26. Purify aRNA (see Basic Protocol 1, steps 28 to 46). The aRNA size should range from 200 to 700 nucleotides.

27. Analyze on Bioanalyzer (see Support Protocol 1) and determine RNA concentration. The before-fragmentation-aRNA concentration is used for calculating the amount hybridized to the Genechip. Example of the aRNA following two-cycle amplification is given in Figure 1B.2.1, lane 4.

Fragment aRNA 28. Fragment the aRNA (see Basic Protocol 1, steps 49 to 52) and analyze the profile. The fragmentation profile should resemble the one-cycle aRNA fragmentation profile. BASIC PROTOCOL 2

Microarray Analysis of Stem Cell Gene Expression

HYBRIDIZING, WASHING, STAINING, AND SCANNING OF THE GENECHIP Affymetrix recommends 15 to 20 µg of fragmented aRNA sample be mixed with hybridization cocktail and hybridized to each Genechip. The hybridization cocktail includes the aRNA, commercially available prokaryotic hybridization controls, bovine serum albumin (BSA), and salmon sperm DNA blocking reagents, 2× hybridization buffer, and water. The cocktail is hybridized 16 hr at 45◦ C, rotating to allow for mixing. On the following day, the wash and stain procedure is performed using the automated Affymetrix

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Fluidics 450 instrument. During this process, Genechips are (1) washed to remove unbound, non-specific hybridization components and (2) incubated with streptavidin fluorescent–conjugate (for binding to bioin-modified-aRNA) allowing for detection of the hybridized target. The Genechips are then scanned using the Affymetrix 3000 Laser Scanner. Signal intensity files are generated, corresponding to the relative levels of gene expression per target or gene. These instructions are modified from the Affymetrix instruction manual available at http://www.affymetrix.com/support/technical/ manual/expression manual.affx.

Materials Genechip eukaryotic hybridization control kit (Affymetrix cat. no. 900299; prokaryotic control samples used for eukaryotic samples) Fragmented ES or EB aRNA (see Basic Protocol 1 or Alternate Protocol 1) Herring sperm DNA (Invitrogen Life Technologies cat. no. 1563-4017) 50 mg/ml acetylated bovine serum albumin (BSA; Invitrogen Life Technologies cat. no. 15561-020) 2× hybridization buffer (Quality Biological cat. no. 351-289-061 or see recipe) Nonstringent wash buffer (Quality Biological cat. no. 351-287-131 or see recipe) 2× MES stain buffer (Quality Biological cat. no. 351-288-101 or see recipe) Streptavidin, R-phycoerythrin streptavidin conjugate (SAPE; Invitrogen Life Technologies cat. no. S-866) Goat anti-streptavidin antibody (Vector Laboratories, cat. no. BA-0500), biotinylated Goat IgG (Sigma-Aldrich, cat. no. I 5256), reagent-grade Stringent wash buffer (Quality Biological cat. no. 351-286-131 or see recipe) 45◦ and 95◦ C heating blocks Affymetrix Genechip hybridization rotating oven Affymetrix Fluidics 450 wash and stain station 1.5- to 1.7-ml nuclease-free tubes Affymetrix Genechip scanner 3000 Genechip operating system software (GCOS) Prepare hybridization cocktail reagents per one Genechip Array 1. Prepare the following master hybridization mix: 21 µg fragmented ES or EB aRNA 5 µl control oligonucleotide B2 (3 nM) 3 µl 20× eukaryotic hybridization controls (bioB, bioC, bioD, cre) 3 µl of 10 mg/ml herring sperm DNA 3 µl of 50 mg/ml BSA 125 µl 2× hybridization buffer Bring to 250 µl with nuclease-free water. 2. Once all components are added, heat the hybridization aRNA cocktail 5 min in a 95◦ C heating block. 3. Heat cocktail 10 min at 45◦ C. 4. Centrifuge 5 min at 13,000 × g, room temperature, to remove insoluble particulates. 5. Remove supernatant and place at room temperature until ready for application to chip (within minutes). The Affymetrix Genechip arrays should be room temperature before using.

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6. For prehybridization, add ∼225 µl of 1× hybridization buffer to each Genechip. Do not fill completely, leave a bubble to allow for mixing of the solution. 7. Rotate 5 min at 45◦ C at ∼60 rpm to mix. If using the recommended hybridization oven, the rotation speed can be entered directly. If another source is used, the speed has to be estimated.

8. Remove the 1× hybridization solution from the Genechip and replace with ∼225 µl of the master hybridization aRNA cocktail. Again, leave a small bubble in the Genechip to allow for mixing of the solution. 9. Place the Genechips with hybridization aRNA cocktail, on rotator. Hybridize 16 hr at 45◦ C, rotating at 60 rpm. 10. Remove the hybridization cocktail and discard. Completely fill Genechip with nonstringent wash buffer (∼250 µl of solution).

Prepare SAPE and antibody for wash and stain procedure Volumes below are for one Genechip. The solutions should be prepared as a master mix; depending on the total number of Genechips to be analyzed, include 5% over volume total to account for pipetting errors. 11. Prepare 1200 µl streptavidin, R-phycoerythrin conjugate (SAPE) as follows:

600 µl 2× MES stain buffer (1× final) 48 µl 50 mg/ml acetylated BSA (2 mg/ml final) 12 µl 1 mg/ml SAPE (10 µg/ml final) 540 µl nuclease-free water. 12. Prepare 600 µl biotin anti-streptavidin antibody solution as follows:

300 µl 2× MES stain buffer (1× final) 24 µl 50 mg/ml acetylated BSA (2 mg/ml final) 6 µl 10 mg/ml normal goat IgG (0.1 mg/ml final) 3.6 µl 0.5 mg/ml biotinylated anti-streptavidin antibody (3 µg/ml final) 266.4 µl nuclease-free water. Prime Fluidics 450 wash and stain station 13. Turn on the Fluidics 450 wash and stain station. 14. Fill buffer reservoirs: add nonstringent wash solution to “A” designated position, stringent wash solution to “B” position, and water to “water” position. 15. Place clean, empty 1.5 to 1.7 ml nuclease-free tubes at positions 1 to 3 on the fluidics station. 16. Open GCOS program on computer. 17. Select Run/Fluidics/Station 1. On access to wash modules, select All Modules. 18. Open Protocol menu and select Prime-450. 19. Select Run. 20. After the priming is done, discard empty tubes from Fluidics station. Microarray Analysis of Stem Cell Gene Expression

Create experimental template 21. From the File menu, select New Experiment. The experimental template contains basic information about the individual Genechips.

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22. Create (1) a sample name, (2) a sample type, (3) a project name, (4) an experiment name (must be unique for each Genehip), and (5) probe-array type used (i.e., probeset genomic library). Optional, scan the Barcode (unique to each Genechip). The New Experiment must be created for each Genechip. Once created, the information becomes accessible in the Fluidics 450 program.

Wash and stain ESC and EB samples 23. Remove the nuclease-free tubes used during the priming protocol. Replace with 600 µl of SAPE in positions 1 and 3, and 600 µl of antibody solution in position 2. 24. Place Genechips in modules. 25. In the GCOS Fluidics protocol, use drop-down menu to designate which Genechip is in a particular module. 26. Select fluidics protocol EukGE-WS2v4 from the drop-down menu. Select Run. Repeat for each module or Genechip. 27. At the end of the run, Fluidics 450 fills each Genechip with nonstringent wash buffer. Inspect to ensure that the Genechip is completely filled and there is no visible debris, dried buffer, etc. on the surface of the Genechip. If necessary, add additional nonstringent wash buffer to completely fill the Genechip, and wipe with 70% alcohol using a lint-free paper towel to clean the outside surface of the Genechip.

Scan chips 28. Turn on the Scanner. Allow at least 10 min to warm up. 29. Choose RUN-Scan. 30. Select Open scanner to open top of scanner, and insert Genechip. 31. Select OK to scan. Results of the scan are generated as DAT/CEL files (CEL file; Fig. 1B.2.2).

Figure 1B.2.2 CEL intensity file. The CEL intensity file for the U133 plus 2 Affymetrix Genechip. Pseudo-color for the graphical display of the relative signal intensities of individual probes on the Genechip. The white fluorescence squares on each square correspond to higher intensity and blue squares correspond to lower intensity.

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32. Select Analysis to analyze the CEL file and generate signal intensities corresponding to individual targets or genes. 33. Select Scaled. Optional: Normalize against a set of housekeeping genes; if this is done, the directions for performing normalization (and the gene lists) are located on the Affymetrix website (designated as Mask files; http://www.affymetrix.com/ products/arrays/specific/hgu133plus.affx). Following analysis, the output files are in the form of Excel-compatible CHP files.

Use report file: quality metrics for Genechip performance 34. From the File menu, select Report to generate the Report file for each Genechip. Examine the following parameters (values for good-quality Genechip hybridizations are provided below in parentheses): (a) Scaling factor: target intensity (500) /average signal intensity (<10) (b) Background: lower 2% signal intensity on Genechip (<120) (c) Noise: pixel-to-pixel variation (<20) (d) Percent genes present: targets with detectable expression (∼40%) (e) 3 /5 ratio GAPDH and actin: assessment of full length transcription (1:5 ratio) (f) Hybridization controls: assessment of hybridization (linear intensities: 1.5 pM BioB, 5 pM BioC, 25 pM BioD, 100 pM Cre). The percent genes present value is a measure of the transcripts detected, which is an indication of transcriptional activity. The value depends on the cell type analyzed. The percent genes present should be fairly consistent between ESC and EB samples (as the number of differentially expressed genes should be comparatively small). Scaling factor values should not vary by more than a factor of 3 between samples being compared. BioB, BioC, BioD, and Cre are prokaryotic hybridization controls added at increasing concentrations to the hybridization cocktail; the resulting signal intensities should be linear. BASIC PROTOCOL 3

Microarray Analysis of Stem Cell Gene Expression

DATA ANALYSIS USING GCOS BATCH ANALYSIS TOOL For each target or gene on the Affymetrix Genechip, there are 10 to 20 of each perfect match (PM) and mismatch (MM) overlapping oligonucleotides or probes, collectively referred to as a probe-set. PMs and MMs are 25-mer oligonucleotides or probes. PM corresponds to the perfect compliment of the target of interest. MM differs from PM by 1 base at the 13th position; designed to represent nonspecific hybridization signal intensity. The Affymetrix GCOS batch analysis tool considers both PM and MM signal intensities to calculate the final intensity of a particular target, and ultimately differential gene expression. PM is not controversial; however, use of MM to determine final signal intensity is controversial. Studies (Li and Wong, 2001; Irizarry et al., 2003) suggest that use of MM leads to variance, which affects reliable selection of differentially expressed candidate targets. As a result, the Affymetrix GCOS batch analysis method is outlined as an optional analysis method. Alternatively, there are a significant number of programs available that disregard MM intensities. These programs are referred to as PM-only analysis methods. As an alternative approach, a PM-only method is outlined in Alternate Protocol 2; also additional Microarray Resources (Table 1B.2.1) useful for data analysis are described. A PM-only method is described, however, programming skills are required for its use. Each of the websites listed in Table 1B.2.1 contain an extensive list of programs allowing for PM-only analysis for determining differential expression, cluster analyses, gene ontology, and pathway analyses as described by many different sources.

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Table 1B.2.1 Websites useful for Analysis of Microarray Datasets

Microarray Resources/Web linksa http://www.cbi.pku.edu.cn/mirror/microarray/soft.html http://genome-www5.stanford.edu/resources/restech.shtml http://biocompare.com/matrix/17648/microarray-analysis-(free-downloads).html http://www.statsci.org/micrarra/ http://fgc.urmc.rochester.edu/data analysis.html http://www.cochranlab.org/BGcourse/Microarray links.htm http://www.usc.edu/hsc/nml/lib-services/bioinformatics/array tool.html http://info.med.yale.edu/wmkeck/dnaarrays/softwarelinks.htm http://abs.cit.nih.gov/ http://nciarray.nci.nih.gov/ a Each website contains links to additional websites for statistical, clustering, pathway, and other software

for analysis of datasets.

A list of different programs are available, most of which are easy to use and free, and nearly all require either the CEL or the CHP files generated by Affymetrix GCOS. As a suggestion, the http://www.cbi.pku.edu.cn/mirror/microarray/soft.html website contains links to user-friendly, step-by-step statistical analysis programs like GEDA from the University of Pittsburg and the Cluster analysis program available from the Micheal Eisen laboratory at Lawrence Berkeley Laboratory. Data analysis software useful for analysis and interpretation of high-throughput microarray data may be available through your university or institute.

Materials Affymetrix GCOS microarray platform GCOS software (available free of charge with the purchase of the Affymetrix microarray platform) 1. Open GCOS software. From Tool-bar, select Shorts, so that the shortcut to Data Sources is displayed. 2. Experiment, DAT, CEL, and Analysis Results (CHP) files should appear. 3. Double click on Analysis Results to display the list of CHP files that are to be compared. 4. From Tool-bar, select Run/Batch Analysis. 5. Drag the ES CHP data to “Input” area. 6. Double click on Baseline to designate EB sample to be used as baseline. Select EB sample from the list that appears. 7. Double click on Output and enter a new file name. 8. Select Analyze from File menu. Output is ESC/EB comparison with the newly designated file name. The output can be reversed by changing the input/baseline to EB/ESC.

9. Refresh CHP files and open the newly generated ESC/EB comparison. ESC/EB CHP comparisons are presented as “nc” (no change), i (increase), d (decrease), or mi (marginally increased) or md (marginally decreased).

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10. Sort the dataset; right click the mouse and designate Sort. Remove nc, mi, and md probe-sets by highlighting nc, mi, and md probe-sets, and selecting Hide from the tool-bar. The i and d differentially expressed probe-sets should remain.

11. Right-click the mouse and sort according to p-value. Probe-sets with values >0.05 are considered less reliable. Hide probe-sets with p values >0.05, by highlighting those samples, and selecting the Hide button from the tool-bar menu. 12. Sort again according to Log ratio to rank probe-sets according to degree of differential expression. The resulting genes represent Selection of candidate gene list. ALTERNATE PROTOCOL 2

DATA ANALYSIS USING MODIFIED PM-ONLY METHOD The approach outlined below is similar in concept to other data analysis programs that consider the PM-only signal intensities extracted from GCOS datasets. The difference between this method and most other PM-only methods, is that this method considers updated corrections to the RefSeq and mRNA databases allowing for accurate target/gene assignments.

Download and install required programs 1. Download and install the most updated BLAST search engine from the NCBI website (http://www.ncbi.nlm.nih.gov/BLAST/download.shtml). 2. Download and install Perl from (http://www.cpan.org). 3. Install the R (http://cran.r-project.org) and BioConductor Package (http://www. bioconductor.org). 4. Install R-Com from (http://cran.r-project.org).

Analyze data 5. Update the Affymetrix Genechip sequence annotation information. a. Download probe level sequences from the Affymetrix website (http://www. affymetrix.com) and format as FASTA sequences. b. Download the most recent release of the Refseq sequences from NCBI (ftp://ftp. ncbi.nih.gov/refseq) and mRNA sequence database from UCSC (http:// hgdownload.cse.ucsc.edu/downloads.html), both in FASTA sequence format. c. Place all sequence files and the executable Blast search files downloaded from NCBI into the same directory. Run “format db” (i.e., database) against either the Refseq or mRNA sequences to generate a searchable Blast database. d. Blast the probe sequences against the Refseq database requiring 100% matching between the probe sequence and Refseq sequence; assign the corresponding annotation to Affymetrix probes. Using this approach, the number of probes correctly assigned to a particular probe-set is numerated.

Microarray Analysis of Stem Cell Gene Expression

e. If none of the probes within the probe-set map to the Refseq, blast the probes against the mRNA sequence database. Assign Genbank identifications to probes mapping to the mRNA database. f. Remove probe-sets that fail to map to either database for further data analysis due to lack of information.

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6. Extract probe level PM information from the ESC and EB hybridization array data sets. a. Using the Bioconductor software package, extract the probe level signal intensities of PM probes from CEL files generated from GCOS.

library(affy); library(matchprobes); library(hgu133plus2probe); d <- ReadAffy(); dmas <- bg.correct(d, “mas”); i.pm <- unlist(indexProbes(dmas, “pm”)); i.pt <- xy2i(hgu133plus2probe$x, hgu133plus2probe$y); index <- match(i.pm, i.pt); seqnc <- hgu133plus2probe$sequence[index]; write.table(cbind(probeNames(dmas), pm(dmas), seqnc), file= “outfile.txt”, sep=“\t”); The file generated above contains all PM information, including those incorrectly assigned.

b. Compare PM information extracted using Bioconductor to the corrected information (derived from the updated Blast; see step 1), using Perl or Bioconductor-R software. The incorrectly assigned probes are removed at this point.

c. For each probe-set, compute the weighted median of the PM intensities using the Tukey Biweight method (Hoaglin et al., 2000; http://www.affymetrix. com/support/technical/whitepapers/sadd whitepaper.pdf) 7. Combine and normalize data. a. For this experiment, SA02 ESC and SA02 EB, and BG01 ESC and BG01 EB samples are hybridized, all as duplicates, for a total of eight Genechips. Process SA02 datasets (i.e., four Genechips) independently of the BG01 datasets (i.e., four Genechips). Generate the gene expression ratios by directly comparing the signal intensities of the ESC/EB samples. b. Perform normalization using Cyclic Lowess accessed via Bioconductor, MEV, or Perl script. Once normalized, generate the gene expression ratios for individual probe-sets of ESC and EB comparison, resulting in four data points for each probe-set for SA02 samples, and four data points for each probe-set for BG01 samples. There are four comparisons, resulting in four data points per probe-set. c. Perform one sample t-test to determine the reliability of the differential ratios from the four replicated values. Probe-set ratios demonstrating p ≤0.05 are determined reliable and considered for further analysis. d. Filter candidate genes according to a fold-change or z-test using standard deviation cutoffs (i.e., 1 or 2 standard deviations).

REAL-TIME PCR VALIDATION A list of differentially expressed candidate genes can be chosen based on reliability of the ‘call’ (i.e., p values <0.05 for increase or decrease designation), degree of differential expression (i.e., ratio of ESC/EB), and consistency of direction of differential expression across replicate samples. Once candidate genes are selected, the Genechip microarray results should be validated. Frequently, this is accomplished by examining expression levels of microarray RNA samples using an alternative method, such as real-time PCR in an effort to demonstrate concordance between the two methods. Current Protocols in Stem Cell Biology

BASIC PROTOCOL 4

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A select number of candidate genes are chosen for real-time PCR validation (see Anticipated Results). Retrieving the precise sequence used for generating the individual probe-sets and plugging it into a primer-generating software is recommended. When using Affymetrix platforms, this sequence is available through Affymetrix’s NetAffxTM Analysis Center (http://www.affymetrix.com/analysis/index.affx). Once the sequence is retrieved, it can be copied and pasted directly into primer programs similar to Primer3 (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3 www.cgi). Additional primergenerating resources and manufacturers are located at http://www.realtimeprimers. org/Links/Links%20Page.html. PCR primers used for this study were manufactured by Integrated DNA Technologies (http://www.idtdna.com/Home/Home.aspx). This method is based on the Ambion MessageAmp II protocol (cat. no. 1791).

Materials First-strand cDNA (see Basic Protocol 1, steps 1 to 9) RNase H (10 U/µl; Ambion cat. no. AM12292) Human glucoronidase (GUS) predeveloped control primers (Applied Biosystems cat. no. 4326320E) Q-PCR primers (Integrated DNA Technologies) 2× AmpliGold PCR master mix (Applied Biosystems cat. no. E08415) 1% (w/v) agarose gel, optional SYBR Green PCR master mix (Applied Biosystems cat. no. 4309155) 37◦ C water bath Applied Biosystems 7900HT real-time PCR system BioAnalyzer DNA Lab-on-a-chip (Agilent cat. no. 5067-1504), optional Optical adhesive covers (Applied Biosystems cat. no. 4311971) Centrifuge (e.g., Eppendorf 5804) Generate cDNA (template for real-time PCR) 1. Generate first-strand cDNA as outlined in Basic Protocol 1, steps 1 to 9. 2. Add 1 µl of RNase H, incubate 30 min at 37◦ C. 3. Add water to 100 µl and collect content at bottom of tube by centrifuging. Store up to 2 years at −20◦ C until ready for real-time PCR.

Test primer specificity 4. Adjust all primer concentrations to 10 µM stock solutions. 5. Perform conventional PCR to examine the specificity of each primer using the following:

1 µl forward primer 1 µl reverse primer 1 µl cDNA template (ESC or EB depending on detection observed in microarray) 10 µl 2× AmpliGold PCR master mix Water to 20 µl. 6. Set up thermal cycler with the following parameters:

Microarray Analysis of Stem Cell Gene Expression

1 cycle: 1 cycle: 1 cycle: 40 cyles:

2 min 10 min 15 sec 1 min

50◦ C 95◦ C 95◦ C 60◦ C.

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7. Examine on a BioAnalyzer DNA Lab-on-a-chip or 1% agarose gel. This method differs from the RNA Nano-chip protocol. The DNA is not heated prior to application to the detection chip and the gel/dye mixture ratio is 500:25 µl. A single amplicon should be generated using the particular cDNA template.

Real-Time PCR Using The Abi 7900Ht System 8. Determine the number of samples (including controls) to be analyzed. Use the 96or 384-well Excel template to outline the experiment. For this study, five template samples (SA02 ES, SA02 EB, BG01 ES, BG01 EB, and water) were analyzed for ten candidate genes and the GUS housekeeping gene, using the SYBR Green detection method as described by Applied Biosystems (http://docs.appliedbiosystems.com/pebiodocs/04310251.pdf).

9. Enter the sample names in the desired order, onto the template.

Prepare master mix 10. Prepare master mixes as follows (volumes listed below are quantities per analysis, not accounting for replicates): (a) master mix per different templates (prepare at least 11×): 10 µl SYBR Green mix 1 µl of ES, EB, or water template 2 µl of water (13 µl total). (b) master mix per primers set (prepare at least 5×): 1 µl forward primer of interest 1 µl reverse primer of interest 5 µl of water (7 µl total). (c) master mix per glucoronidase control (prepare at least 5×): 1 µl glucoronidase mix 6 µl of water (7 µl total). The final reaction mixture is 20 µl containing two basic components: (1) template with PCR reagents and (2) primers. The objective here is to analyze each of the five templates for expression of the corresponding eleven transcripts (i.e., as determined using the eleven primers). There are five templates SA02 ESC, SA02 EB, BG01 ESC, BG01 EB, and water (prepared for a total volume of 13 µl; there are eleven primers including ten differentially expressed genes of interest and the GUS control (prepared for a total volume of 7 µl). For each template or primer, individual master mixes are prepared.

Load 96-well microtiter plate 11. Examine each of the templates (five total) for the presence of the eleven genes of interest. Dispense 13 µl of the template and 7 µl of the primer mixture into the designated microtiter well. 12. Cover the plate with optical adhesive cover. Press to remove wrinkles. 13. Centrifuge the plate 1 min at 1500 × g, room temperature.

Use ABI 7900HT Sequence Detector 14. Open the SDS 7900 program. 15. From the menu, go to File and designate New. A new document window will appear.

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16. From this window access:

(a) Assay: allowing for allelic discrimination or quantitative analysis (b) Container: 96- or 384-well plate (c) Template: blank template. 17. For expression analyses, designate Quantitative Analysis. 18. For the plate format, use the drop-down menu to designate 96- or 384-well plate. 19. For the Template, always designate Blank Template. 20. Click OK (the window should disappear). 21. Click on Tools at the top menu. A Detector Manager window should appear.

22. At the bottom of this window click on File, then New. Add Detector Window window should appear.

23. Check that the following appear:

(a) Name: designate the gene and detector here (b) Description: optional; for reference (c) Reporter: designate SYBR Green (d) Quencher: non-fluorescent. Click OK (the detector will be added to the Detector Manager List). 24. Click on Copy to plate document (a new screen will appear behind the Detector Manager). 25. Close the Detector Manager screen by clicking the Done button. A plate document is now ready to be created.

Create a plate document 26. Three panels should appear. (a) The “Untitled template” (panel at the top left) (b) The “Excel version of the template” (noting position of the samples; bottom left) (c) The “Set-up/Instrument” screen (top, right-hand corner; bottom half is the “Instrument control panel”—for open/close of the instrument). 27. Using the 96- (or 384-) well template, enter the sample names per well. 28. If sample 1 is placed in position A1 (and A2, for duplication), highlight A1/A2 (by dragging the mouse over wells A1 and A2). In the Set-up Screen/Sample name, the word “Mixed” should appear. Click on Mixed. You should replace this with a more specific name for the sample to be analyzed. Continue until each well is labeled. 29. Highlight all samples. Go to set-up and click on the box next to SYBR Green detector. This step designates the reporter used to analyze the samples.

Microarray Analysis of Stem Cell Gene Expression

Specify instrument set-up conditions 30. Click on Instrument (next to Set-up). 31. Select Standard mode. Thermal cycler profile/Auto Increment/Ramp Plate/Data Collection should appear.

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32. Designate the Thermal cycle conditions as follows: 1 cycle: 1 cycle: 1 cycle: 40 cycles:

2 min 10 min 15 sec 30 sec

50◦ C 95◦ C 95◦ C 60◦ C.

33. Enter sample volume: 20 µl. 34. Select Connect. StartStopDisconnectOpen/close should appear.

35. Push the Open/close button to open the 7900HT. 36. Make sure that the proper block and instrument trays are inserted, i.e., 96- (or 384-) well block with corresponding instrument tray. 37. Place the plate onto the instrument tray. Make sure that the sample plate is covered and has been centrifuged. Push Open/close again to close the chamber. 38. Push Start to start the instrument.

Data presentation 39. After a successful run, click Analysis or the green icon button (arrowhead; just below Analysis). Right panel should expand to display Set-up, Instrument, Results, and Dissociation Curve.

40. Click on Results. Amplification plot should appear.

41. Adjust the baseline. 42. From the menu, select File/export/results.

Calculate relative gene expression levels 43. Calculate the relative expression level of targets on the delta comparative threshold (
Ct) as suggested by PE Applied Biosystems. Relative ratio of expression = 2–[
Ct 1–
Ct 2] . Delta Ct 1 (
Ct1) is the difference between the Ct value of a particular gene in normal reference sample minus the Ct value in the hESC sample. The
Ct2 is the difference between the Ct value for the beta-glucoronidase (GUS) control gene in normal reference minus the Ct value in the hESC sample. A positive value represents up-regulation and a negative value represents down-regulation with respect to ES samples. The
Ct2 for GUS control has been determined experimentally to be ∼0.5.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Hybridization buffer, 2× For a 50-ml solution, add: 8.3 ml 12× MES stain buffer (see recipe; 100 mM final) 17.7 ml 5 M NaCl (1 M final) 4.0 ml 0.5 M EDTA (20 mM final) 0.1 ml 10% Tween-20 (0.01% v/v final) 19.9 ml water Store up to 1 year at 4◦ C and shield from light

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MES stain buffer, 12× For a 1-liter solution, add: 70.4 g MES free-acid monohydrate (1.22 M final) 193.3 g MES sodium salt (0.89 M final) 800 ml molecular-grade water Mix and adjust volume to 1 liter pH should be between 6.5 and 6.7 Filter through a 0.2-µm filter For a 2× MES stain buffer For 250 ml solution, add: 41.7 ml 12× MES stain buffer (100 mM final) 92.5 ml 5 M NaCl (1 M final) 2.5 ml 10% Tween 20 (0.05% v/v final) 113.3 ml nuclease-free water Filter through a 0.2-µm filter Store up to 1 year at 4◦ C and shield from light Nonstringent wash buffer For 1 liter, add: 300 ml 20× SSPE (6× final) 1.0 ml 10% Tween 20 (0.01% v/v final) 699 ml water Filter through a 0.2-µm filter Store up to 1 year at 4◦ C Stringent wash buffer For 1 liter, add: 83.3 ml 12× MES stain buffer (see recipe; 100 mM final) 5.2 ml 5 M NaCl (0.1 M final) 1.0 ml 10% Tween 20 (0.01% v/v final) Filter through a 0.2-µm filter Store up to 1 year at 4◦ C and shield from light COMMENTARY Background Information

Microarray Analysis of Stem Cell Gene Expression

Stem cells are pluripotent cell types, which have the ability to self-renew and live for long periods of time (Shamblott et al., 1998, 2001; Thomson et al., 1998), implicating them in repair and revitalization of normal tissues. In addition to their role in normal tissues, they are thought to be intricately linked to many different disease states. Studies show that because stem cells are pluripotent they can replenish organs ravaged by diseases like Alzheimer, Parkinson, or any other disease requiring tissue regeneration (Suhr and Gage, 1993, 1999; Sell, 2001; Ginsberg et al., 2006; Miller and Federoff, 2006). Stem cells are also implicated in cancers. Current thinking is that stem cells are the progenitor cancer cell (Dontu et al., 2003a,b; Tatematsu et al., 2003; Dontu et al., 2005) existing in organs in the early stages of cancer, as cancer stem cells. This is supported

by the theory that stem-like cells are the only cells known to live for long time periods, as is required to accumulate mutations necessary for tumorigenesis. ESC and the differentiated EB cell types are characterized in an attempt to better understand the biology of stem cells, so that ultimately their utility in tissue regeneration and possible involvements in cancers can be determined. Much of the data characterizing stem cells comes from analyses using the RNA expression microarray platforms, which began with an analysis using cDNA microarrays (Thomson et al., 1998; Kelly and Rizzino, 2000). In addition to the microarray platform, other high-throughput gene expression methods have been performed for characterization of stem-related cell types, each with advantages and disadvantages (Robson, 2004). More common methods are differential

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display (DD; Hollnagel et al., 1999), serial analysis of gene expression (SAGE; Richards et al., 2006) and massively parallel signature sequencing (MPSS; Brenner et al., 2000; Reinartz et al., 2002; Brandenberger et al., 2004; Bhattacharya et al., 2005). Differential display is a PCR-based method that uses short non-specific primers for amplification, followed by high-resolution gel analysis to detect differentially represented transcripts. In SAGE, double-stranded cDNA is generated, immobilized, tagged, and PCR amplified; the number of tags present being a measure of the transcript copies. In MPSS, a 17-base sequence is generated at a specific site upstream of the poly A tail, which serves as a signature for mRNA; millions of transcripts are sequenced and enumerated, and compared between samples as a measure of differential transcript levels. Unlike the microarray platform, DD, SAGE, and MPSS are considered unbiased methods because nearly all transcripts are thought to be captured and subsequently analyzed. For this reason, these methods are considered more conducive to high-thoughput novel ESC gene discovery (Robson, 2004). Even with its disadvantage, microarrays have become the most popular of platforms, as the aforementioned methods can be expensive, lengthy, and/or technically challenging. Over the past few years, there has been a tremendous accumulation of data in stem cell biology resulting in the discovery of stemrelated genes and pathways. For human stem cells, NANOG, POU5F1/OCT4, and SOX2 are nearly always identified (Bhattacharya et al., 2004; Skottman et al., 2005; Yang et al., 2005), and by functional analysis, demonstrated to be essential in maintaining the stem cell state (Tsuji et al., 2006; Wu et al., 2006; Zhang et al., 2006a). A recent and very convincing study by Takahasi et al. (Takahashi and Yamanaka, 2006) showed SOX2, OCT4, KLF4, and MYC were sufficient for reprogramming mouse fibroblasts to stem cell state, demonstrating the importance of these genes in effecting stemness, in mice. For many of these studies, large numbers of genes are identified as stem-related, with fewer common between studies. This could be due to differences in the biology or physiological state of the various cell types, and technical variations between experiments and laboratories. To begin to address these problems, a number of different options have been proposed. The Microarray Quality Control Consortium (MAQC) is a group of 137 investigators and

51 different organizations (Shi et al., 2005) whose goal is to examine the reproducibility of the high-density multi-platform microarrays and make suggestions on improving reproducibly within and between laboratories. The group proposed and is studying the utility of universal amplification and hybridization RNA controls (Tong et al., 2006). Similarly, universal reference hybridization controls have been suggested for use in the two-color array platforms (Novoradovskaya et al., 2004). The MGED group recommends that Minimum Information About Microarray (MIAME; Brazma et al., 2001) be implemented to document the experimental design, samples used, sample preparation, hybridization, data measurement, data analysis, and assay design so that investigators can replicate each other’s experiments. MIAME guidelines consider both the biology of the samples and technical issues related to the protocol. Use of the various controls and MIAME implementation should greatly improve reproducibly between studies, as array platforms themselves appear somewhat robust. Studies show that when the same biological samples are examined on different platforms and in different laboratories, there is remarkable concordance between experimental results (Canales et al., 2006; Shi et al., 2006). This suggests that the platforms themselves are fairly reliable. Discordance between studies are possibly related to experimental or technical variation between laboratories and/or physiological variation between biological samples (i.e., RNA expression levels related to cell cycle differences, cell culture conditions, or stages of cellular differentiation). The microarray platform is very useful as the first approach for global analysis of ESC samples. It should not be considered the final analysis platform. For reliability, technical and biological replicates should be hybridized, and the data validated and analyzed using a number of different analytical methods. Probably the most time-consuming part of the entire process is analyzing the data, which could take weeks to months. Once a final candidate gene list is generated, gene expression levels are validated using real-time PCR. Data can be further analyzed using different analysis softwares, such as clustering, gene ontology, and pathway tools in an effort to identify patterns associated with the differentially expressed genes. Commonly, candidate gene lists contain many genes, so additional analysis tools are necessary to make sense of it all. The GeneSet Enrichment Analysis (GSEA; http://www.broad.

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mit.edu/gsea) is a particularly useful tool for identifying genes common within and across experimental datasets. It is unlikely that a single gene is responsible for the very complex processes that define the ESC or EB cell types, therefore, it is important that programs useful for identifying trends or patterns be used for examination of the candidate gene lists. Once this is done, biological function has to be validated using studies that allow for either re-introduction (if down-regulation) or inhibition (if up-regulation) of the gene of interest. Although not always successful for a number of different reasons, immunohistochemistry can be performed to demonstrate concordance between transcript and protein levels. In situ methods are key for determining localization within the cell, which is an initial step in characterizing the function of the genes of interest. There are basically three types of microarray platforms utilized for the analysis of RNA transcript levels, all of which can be used for analysis of transcript levels in stem cell samples. They are manufactured by immobilizing short oligonucleotides (25 to 30 bases), long oligonucleotides (60 to 70 bases), or copyDNA (cDNA clone based) of varying lengths, on various types of substrates (Barrett and Kawasaki, 2003), all of which can accommodate the full genome. Oligonucleotide arrays may be synthesized in situ by photolithographic means (Fodor et al., 1991) or by inkjet mechanisms (Hughes et al., 2001). Other oligonucleotide and cDNA clone arrays are manufactured using robotic pin printers or microfluidic instruments (Hughes et al., 2001; Situma et al., 2006). For nearly all platforms, sense molecules are immobilized onto a surface, so that they are available for hybridization to the antisense complementary sequence (i.e., hESC and EB biological sample). The antisense material is first-stand cDNA generated using reverse transcriptase or antisenseRNA (aRNA) amplified to obtain higher concentrations of the samples’ messenger RNA species. Detection of hybridization to microarrays has developed as either one- or two-color fluorophore systems. Two-color arrays require labeling of the control sample such as ESC with one fluorophore (usually Cy3) and the test sample such as EB with another fluorophore (usually Cy5), or vice versa. Samples are mixed prior to hybridization, resulting in a competitive hybridization situation between the test and control sample. After hybridization, analysis of the ratios of Cy3/Cy5 levels in the hybrids is compared to determine differential transcript levels between samples. For

single-color arrays, an individual ESC or EB sample is labeled and hybridized per array; differential transcript levels are determined by comparing signal intensities between arrays. Each of the three types of micorarray platforms described above can be generated as either high-density (i.e., containing sequences representing the entire transcriptome) or lower-density microarray platforms. High-density micorarrays are nearly always commercially made, while lower-density microarrays are generated either as a custom commercial microarray or by the investigator. The lower-density arrays, frequently termed focused microarrays, are less complex and usually contain targets or genes representing a particular pathway or chromosome, depending on the needs of the investigator. Lower-density microarrays generally contain <5000 genes. These studies allow for RNA expression analysis of smaller numbers of genes, without the complexities associated with high-density microarrays. As an example, focused arrays for stem cell studies would contain ESC- and/or EB-associated genes, and genes involved in Notch and Wnt pathways, just to name a few. Generally two-fold less RNA is hybridized to the focused arrays than the high-density arrays, therefore, fewer cells are needed. Cell number and RNA concentration requirements for the high-density microarray platforms are discussed below and in Critical Parameters. For this unit, the analysis of ESC or EB samples using a high-density microarray platform is described and results are compared to those previously published. Many microarray analyses studies have been performed, using each of the various platform designs. Even though large datasets are generated, characterizing ESC and/or EB samples, some genes appear common between analyses (Brandenberger et al., 2004; Richards et al., 2004; Skottman et al., 2005). Investigators have concluded that when sequence-to-sequence, as opposed to gene-to-gene, comparisons are performed, results between studies are to some degree comparable (Mecham et al., 2004; Shi et al., 2005, 2006). The Affymetrix (http://www.Affymetrix. com) microarray platform is currently the most widely used commercial platform for RNA expression analysis (as determined by the number of PubMed citations). Other vendors include Agilent (http://www.Agilent. com), Applied Biosystems (http:// www.Appliedbiosystems.com), Nimblegen (http://www.Nimblegen.com), and Illumina (http://www.Illumina.com), just to name a few

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of the larger companies. Early on, Affymetrix was the only commercial vendor offering a platform that allowed for analysis of the entire transcriptome; for this reason, the authors used Affymetrix for their studies and have continued to include Affymetrix analyses for direct comparisons to previous experimental datasets. Requirements and protocols for performing the Affymetrix procedure are detailed in the protocols, much of which are applicable to use with other expression microarray platforms. Other than Affymetrix, the authors have only examined the Illumina microarray platform, which is very different from most. Unique to the Illumina platform are (1) sense 50-mer oligonucleotides are conjugated to magnetic beads, (2) sense molecules, which correspond to a particular target or gene are represented up to 30 times, allowing for statistically reliable calculation of the target or gene signal intensities, eliminating the need for replicate hybridizations, and (3) unlike other platforms, nanogram quantities of aRNA (ESC or EB) can be applied for hybridization. While other platforms require microgram quantities of hybridization material, as little as 750 ng of aRNA are sufficient for hybridization to the Illumina microarray platform. This can be particularly advantageous when small amounts of sample or small numbers of cells are available for RNA extraction, which frequently is the case when studying stem cell preparations. In excess of 750 ng of aRNA can be generated from as few as a thousand ESC or EB cells. These calculations are based on an estimate of 10 pg of total RNA per cell, 1000 cells, and 10 ng of total RNA purified from the cells. For Illumina and a number of other microarray platforms, total RNA is then amplified to generate amounts necessary for hybridization. In this case, 10 ng of total RNA can be amplified to generate up to 1.0 µg of aRNA. An RNA amplification method similar to the one used for an Illumina platform is described in Alternate Protocol 1. For more information describing the Illumina procedure, documents are available for download at http://www. illumina.com/pages.ilmm?ID=208. Unlike earlier years, Illumina and most other commercial vendors now supply a variety of different platform options, including custom platforms and those allowing for analysis of the entire transcriptome. If the number of cells and subsequent amount of RNA is less of an issue, then the choice of microarray platform becomes one of personal preference.

Shortcomings of the microarray platforms Lack of standard protocols Although gene expression platforms have proven to be instrumental in the highthroughput analysis of transcript expression levels and characterization of ESC and EB samples, there is still room for improvement in the protocol. Considering the volume of data analyzed over the past 10 years alone using microarrays, one would expect significantly more insights into the biology of ESC and EB cell types. There are possible explanations for the relatively slow progress and some inconsistencies between results generated between studies. MAQC recommends that reference spike-in standards be used during amplification and as hybridization controls (Tong et al., 2006) for all microarray experiments; MIAME requires that experimental detail accompany each dataset (Brazma et al., 2001), so that other investigators can more easily reproduce the data. It is likely that compliance with MAQC and MIAME will lead to further improvements in conformity of the experimental results. Microarray is not an ideal platform for novel gene discovery The Affymetrix U133 plus 2 Genechip contains over 54,000 probe-sets including Refseq and ESTs (http://www.affymetrix.com/ support/technical/datasheets/human datasheet. pdf). Similar to other platforms, it has been useful for identification of the genes and pathways involved in differentiation of ESC to EB (Zhan et al., 2005; Player et al., 2006; Zhang et al., 2006a,b). Even though there are many ESTs on the microarray, it appears to be less suitable for novel gene discovery compared to SAGE and MPSS. The SAGE and MPSS protocols are shot-gun like approaches. SAGE and MPSS are designed such that mRNA species are immobilized and subsequently interrogated, regardless of the sequence, leading to analysis of hundreds of thousands to millions of both characterized and uncharacterized transcripts. This is not to suggest that more is better, but that shot-gun like approaches that capture, and subsequently analyze larger numbers of transcripts might be more conducive to discovering novel differentially expressed genes. Issues with annotation A working draft of the human genome was made available to the public after June 2000. Even though the bulk of the sequencing efforts are complete, corrections to the

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genome assembly are updated on a quarterly basis (ftp://ftp.ncbi.nih.gov/refseq/ and http://hgdownload.cse.ucsc.edu/goldenPath/ hg18/bigZips/). Most corrections are in annotation or labeling of the relevant genomic regions. Many of the commercial microarray platforms are designed using outdated databases, hence outdated annotation data. This can be a problem when characterizing differentially expressed targets or genes, leading to inaccuracies in target identity. The PM-only analysis method described in this unit, utilizes the most current versions of the sequencing databases to avoid such problems. Differential expression levels A common practice is to rank differentially expressed genes according to degree of differential expression; the most highly differentially expressed genes are then used for further examination. This is an acceptable approach for the initial characterization. However, care must be taken to not overlook targets demonstrating low levels of differential expression (i.e., less than two-fold); instead, consider all targets that demonstrate reliable, consistent patterns of differential expression.

Critical Parameters It is very important that quality metric parameters be established from the beginning, and not compromised for the duration of the study. This includes (1) standardizing the total input RNA concentration to be used for the study and assessing RNA quality, (2) following hybridization, establishing acceptable parameters for the quality metrics listed on the GCOS Report file, and (3) thoroughly analyzing the data using a variety of different methods.

Microarray Analysis of Stem Cell Gene Expression

Input RNA concentration and quality Input RNA concentrations for ES and EB samples, should be the same, so as to preserve (as closely as possible) the mRNA representation of the samples. Input concentrations are also important because it determines if one- or two-cycle aRNA amplification is necessary to achieve adequate concentrations for hybridization to the Genechips. If using ESC and EB RNA concentrations of 1 µg to 200 ng, then one cycle of amplification is sufficient. Assuming a conservative estimate of 10 pg total RNA per cell, this represents amounts obtainable from 105 to 2 × 104 actively dividing cells. If 1 µg of RNA is chosen, then the same input concentration has to be used for all cell preparations, for the duration of the study. If RNA amounts are limited (i.e., 200 ng down to 10 ng),

then consider two rounds of amplification to accumulate enough material for performing technical replicates (i.e., ∼40 µg to be split between two Genechips). Technical replicates are defined as the same aRNA sample analyzed on two different genechips; technical replicates are performed to assess reproducibility of the technique. For RNA concentrations <10 ng, the investigator might consider some other method of analysis other than global expression analysis. Total RNA concentrations down to single cell levels (i.e., 10 to 50 pg) have been amplified using Epicentre’s Target Amp, Genisphere’s Ramp-Up, Qiagen’s Sensiscript, and Superarray’s PicoAmp (A.. Player, unpub. observ.), and later examined using microarray (Tietjen et al., 2003; Hartmann and Klein, 2006), allowing for analysis of differential expression. However, currently, there are significant technical limitations to performing these experiments, so they are not widely adopted. In addition to technical errors, with low concentrations, one risks generating nonspecific, biased amplification products (Peano et al., 2006; A. Player, unpub. observ.), which in most cases are not suitable or adequate for full genome analysis using the microarray platforms. Total RNA should be examined and shown to be of good quality. As an example, A260/280 ratios of 1.85 to 2.1, RNA integrity numbers of 10.0 to 7.0, or 28S to 18S ratios of 2.0 to 1.4 are indicative of good quality RNA. For the first round of RNA amplification, expect at least 1000-fold amplification (calculated based on starting mRNA concentration). Inefficient, less than expected amplification is indicative of poor RNA quality, technical errors, or impurities in the samples. All samples should fall within the established ranges substantiating their overall similarity. Hybridization Technical replicates should always be performed, as they are as important as biological replicates. Technical replicates are also performed to determine reliability of the signal intensities for each gene. The authors routinely compare the signal intensities between technical replicates, only extracting those performing similarly on each Genechip; this is determined by Perl script analysis. Following hybridization and analysis of the Affymetrix Genechip, the Report file is generated for each ESC and EB Genechip. The Report contains the following quality metrics: scaling factor, background, percent genes present, prokaryotic control values, and 3 /5 control values. All

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Genechips should have comparable quality metrics. Scaling factors are calculated based on Genechip signal intensity and should not vary by more than a factor of three between Genechips; larger variance indicates significant differences in signal intensities between Genechips. Analyze and validate results Data should be analyzed using statistical methods, and filtered to generate a reliable, final candidate gene list. However, it is important to note that the most highly differentially expressed genes might not always be the most biologically relevant. Data should also be analyzed using clustering, gene ontology, and other analysis tools. These programs are particularly useful because they can demonstrate relatedness or similarities associated with genes. It is more likely that particular pathways are important as opposed to individual genes in defining ESC and EB. Generally, these programs are very simple to use and do not require programming skills. Investigators can personally perform many of the analyses, allowing for direct, consistent contact with their experimental data. RNA expression data must also be validated using other platforms; often, real-time PCR is performed first using the same biological RNA samples. Candidate gene expression can then be substantiated by analyzing additional clinical samples, immunohistochemistry protein analysis, and functional biological assays such as transfection studies.

Troubleshooting This is a very detailed experimental application, involving a large number of processes. If quality control metrics are established at various points for each procedure, then it becomes easier to identify the problem steps.

Anticipated Results To ensure the selection of reliable candidate genes, strict quality control metrics are applied throughout the entire experimental procedure, and targets are chosen based on reliability of differential expression ratios (i.e., p values <0.05 of increased or decreased targets), consistency of direction of differential expression (i.e., up or down regulation in ESC/EB), and lastly, degree of differential expression. All datasets are analyzed using the PM-only analysis method. There are >54,000 probe-sets on the U133 plus 2 Genechip. The above-mentioned selection criteria are necessary to generate a working list of candidate genes of ∼1500 for SA02 or BG01. SA02 ESC RNA was compared to SA02 EB; BG01 ESC RNA was compared to BG01 EB. Then the SA02 candidate list was compared to the BG01, generating 563 genes common between the two sample types using a PERL script. The most reliable, highly differentially expressed are listed in Table 1B.2.2. The final candidate gene list contained SOX2, NANOG, POU5F1, TDGF1, LIN28, DNMt3B, CD 44, and CD 24, which were previously identified by Skottman et al. (2005) and others (Sato et al., 2003; Abeyta et al., 2004; Bhattacharya et al., 2004). For the current study, ten genes were selected for real-time PCR validation. The genes included SOX2, NANOG, POU5F1, DNMt3B, and CD24 (as identified by other studies) and five additional genes, including DPPA4, HOXC6, MAF, CXCR7, and MEIS1. Except for HOXC6 in the SA02 sample, results were consistent with microarray expression results (Fig. 1B.2.3). Primer sequences are given in Table 1B.2.3.

Figure 1B.2.3 Real-time PCR analysis compared to microarray results. Ten genes were chosen for validation using the real-time PCR protocol, designated by gene symbols along the x-axis. For each of the ten gene of interest, real-time PCR results using SA02 (grey square) and BG01 (red square), compared to Genechip microarray results using SA02 (yellow square) and BG01 (blue square) were mapped to demonstrate concordance. Real-time PCR (
Ct: comparative threshold, ESC/EB) and microarray (log2 ratio ESC/EB) are plotted versus gene symbols.

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Table 1B.2.2 ES/EB Differentially Expressed Candidate Genes

Ratio ES/EB

Gene symbol

Accession number Definition

-6.5

FRZB

NM 001463

frizzled-related protein

-5.6

PRSS35

NM 153362

protease, serine, 35

-5.6

PITX2

NM 153426

paired-like homeodomain transcription factor 2

-5.5

COBLL1

NM 014900

COBL-like 1

-5.4

IGFBP5

NM 000599

insulin-like growth factor binding protein 5

-5.2

SNAI2

NM 003068

snail homolog 2

-5.1

HOXC6

NM 004503

homeobox C6

-5.1

HOXB3

NM 002146

homeobox B3

-4.8

CD44

NM 001001389

CD44 molecule

-4.8

CXCR7

NM 020311

chemokine (C-X-C motif) receptor 7

-4.8

CDH11

NM 001797

cadherin 11, type 2, OB-cadherin

-4.7

CEBPD

NM 005195

CCAAT/enhancer binding protein

-4.5

MEIS2

NM 020149

Meis1, myeloid ecotropic viral integration site 1

-4.1

HOXB5, B7, B8 NM 004502

homeobox B8

-3.9

FLRT2

NM 013231

fibronectin leucine rich transmembrane protein 2

-3.9

KCTD12

NM 138444

potassium channel tetramerisation domain

-3.8

GATA6

NM 005257

GATA binding protein 6

-3.5

LMCD1

NM 014583

LIM and cysteine-rich domains 1

-3.5

RAI1

NM 030665

retinoic acid induced 1

-3.5

GAS1

NM 002048

growth arrest-specific 1

-3.5

DKK1

NM 012242

dickkopf homolog 1

-3.4

CDC42EP5

NM 145057

CDC42 effector protein

-3.4

RUNX2

NM 001024630

runt-related transcription factor 2

-3.3

PDZRN3

XM 942686

PDZ domain containing RING finger 3

-3.3

KDELR3

NM 006855

KDEL (Lys-Asp-Glu-Leu)

-3.1

CRISPLD2

NM 031476

cysteine-rich secretory protein LCCL domain

-3.1

MAF

NM 001031804

v-maf musculoaponeurotic fibrosarcoma secreted protein, acidic, cysteine-rich

-3.0

SPARC

NM 003118

(osteonectin)

-2.9

TGFB1I1

NM 015927

transforming growth factor beta 1 induced continued

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Table 1B.2.2 ES/EB Differentially Expressed Candidate Genes, continued

Ratio ES/EB

Gene symbol

Accession number Definition

-2.8

HOXB4

NM 024015

homeobox B4

-2.8

HOXA13

NM 000522

homeobox A13

-2.8

HOXB2

NM 002145

homeobox B2

-2.7

PDGFRA

NM 006206

platelet-derived growth factor receptor

-2.6

EDNRB

NM 000115

endothelin receptor type B

-2.4

HOXC4

NM 014620

homeobox C4

-2.4

ABAT

NM 000663

4-aminobutyrate aminotransferase

-2.3

AQP3

NM 004925

aquaporin 3 (Gill blood group)

-2.3

TWIST1

NM 000474

twist homolog 1 (acrocephalosyndactyly 3)

-2.2

BMPR2

NM 001204

bone morphogenetic protein receptor

2.2

HESX1

NM 003865

homeobox, ES cell expressed 1

2.2

CEBPZ

NM 005760

CCAAT/enhancer binding protein zeta

2.3

CTSC

NM 001814

cathepsin C

2.5

TERF1

NM 003218

telomeric repeat binding factor 1

2.5

HMGB3

NM 005342

high-mobility group box 3

2.7

CXCL5

NM 002994

chemokine (C-X-C motif) ligand 5

2.8

AASS

NM 005763

aminoadipate-semialdehyde synthase

2.9

JARID2

NM 004973

jumonji, AT rich interactive domain 2

2.9

MAL2

NM 052886

mal, T-cell differentiation protein 2

2.9

FGF2

NM 002006

fibroblast growth factor 2 (basic)

3.1

RABGAP1L

NM 014857

RAB GTPase activating protein 1-like

3.1

KBTBD8

NM 032505

kelch repeat and BTB (POZ) domain

3.1

KCNG3

NM 133329

potassium voltage-gated channel, subfamily G

3.3

SP8

NM 198956

Sp8 transcription factor

3.4

HEY2

NM 012259

hairy/enhancer-of-split related with YRPW

3.4

SOX2

NM 003106

SRY (sex determining region Y)-box 2

3.5

GRB14

NM 004490

growth factor receptor-bound protein 14

3.6

ITGB1BP3

NM 170678

integrin beta 1 binding protein 3

3.6

CD24

XM 942817

CD24 molecule

3.6

DPPA4

NM 018189

developmental pluripotency associated 4

3.6

FRAT2

NM 012083

frequently rearranged in advanced T-cell

3.6

FGF2 NUDT6

NM 198041

nudix (nucleoside diphosphate linked moiety X)

3.6

PMAIP1

NM 021127

phorbol-12-myristate-13-acetateinduced protein continued Embryonic and Extraembryonic Stem Cells

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Table 1B.2.2 ES/EB Differentially Expressed Candidate Genes, continued

Ratio ES/EB

Gene symbol

Accession number Definition

3.9

IRX2

NM 033267

iroquois homeobox protein 2

4.1

KIFC2

NM 145754

kinesin family member C2

4.4

HAS3

NM 005329

hyaluronan synthase 3

4.5

DNMT3B

NM 175850

DNA (cytosine-5-)-methyltransferase

4.6

VASH2

NM 024749

asohibin 2

4.8

GABRB3

NM 021912

gamma-aminobutyric acid

4.9

N-PAC CXCL5

NM 032569

chemokine (C-X-C motif) ligand 5

4.9

CLDN6

NM 021195

claudin 6

5.0

DPPA2

NM 138815

developmental pluripotency associated 2

5.4

LIN28

NM 024674

lin-28 homolog

5.6

LEFTY1

NM 020997

left-right determination factor 1

5.6

POU5F1

XM 930506

POU domain, class 5

5.9

NANOG

NM 024865

Nanog homeobox

6.1

GAL

NM 015973

galanin

6.4

TDGF3 TDGF1

NM 003212

teratocarcinoma-derived growth factor

6.4

NTS

NM 006183

neurotensin

Table 1B.2.3 Primer Sequences of Genes Analyzed Using Real-Time PCR

Gene symbol

Left primer sequence

Right primer sequence

POU5F1

ggaaagagaaagcgaaccag

gccggttacagaaccacact

DPPA

gcactggaaggagtggaaga

ttctgaccaccaacaaccaa

NANOG

tgggaggctttgcttattttt

tgtcattacgatgcagcaaa

DNMT3

ctcagaggcagtgacagcag

tgtctgaattcccgttctcc

SOX2

accagctcgcagacctacat

tggagtgggaggaagaggta

CD24

tgcctcgacacacataaacc

tcgatctgtttgttcccatgt

HOXC6

tctgctccagggcctcag

ccataatgacccccaggatt

MAF

ttgggactgaattgcactaaga

aaatccagttctgcaccaca

CXCR7

gggaggcatagtgctgacat

ccggtacaaaacaccacaca

MEIS1

aatgccaatgtcagcatcaa

tggaactccacagtcattgc

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Time Considerations Once total RNA is purified, it requires the following (1) 3 days are required for onecycle amplification, examination, and quantification of the aRNA product. If two-cycle amplification is to be done, it takes an additional 1.5 days. (2) An additional 2 days are required to hybridize and perform the wash and stain Genechips protocol. (3) The most time-consuming part is data analysis, which requires weeks to months, as thorough analysis of the data must be done to identify candidate differentially expressed genes present per stem cell samples. (4) Candidate gene lists are generated and results must be validated using real- time PCR. This takes an additional 1 week, which includes generating the cDNA (1/2 day), designing, purchasing, and testing primers. The real-time PCR can be performed using 96- or 394-well plates, which allow analysis of up to 96 or 394 samples, respectively.

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Microarray Analysis of Stem Cell Gene Expression

Lander, E.S., Linton, L.M., Birren, B., Nusbaum, C., Zody, M.C., Baldwin, J., Devon, K., Dewar, K., Doyle, M., FitzHugh, W., Funke, R., Gage, D., Harris, K., Heaford, A., Howland, J., Kann, L., Lehoczky, J., LeVine, R., McEwan, P., McKernan, K., Meldrim, J., Mesirov, J.P., Miranda, C., Morris, W., Naylor, J., Raymond, C., Rosetti, M., Santos, R., Sheridan, A., Sougnez, C., Stange-Thomann, N., Stojanovic, N., Subramanian, A., Wyman, D., Rogers, J., Sulston, J., Ainscough, R., Beck, S., Bentley, D., Burton, J., Clee, C., Carter, N., Coulson, A., Deadman, R., Deloukas, P., Dunham, A.,

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Current Protocols in Stem Cell Biology

Li, J., Spletter, M.L., and Johnson, J.A. 2005. Dissecting tBHQ-induced ARE-driven gene expression through long and short oligonucleotide arrays. Physiol. Genomics 21:43-58. Liu, Y., Shin, S., Zeng, X., Zhan, M., Gonzalez, R., Mueller, F.J., Schwartz, C.M., Xue, H., Li, H., Baker, S.C., Chudin, E., Barker, D.L., McDaniel, T.K., Oeser, S., Loring, J.F., Mattson, M.P., and Rao, M.S. 2006. Genome wide profiling of human embryonic stem cells (hESCs), their derivatives and embryonal carcinoma cells to develop base profiles of U.S. Federal government approved hESC lines. BMC Dev. Biol. 6:20. Loring, J.F. and Rao, M.S. 2006. Establishing standards for the characterization of human embryonic stem cell lines. Stem Cells 24:145-150. Mecham, B.H., Klus, G.T., Strovel, J., Augustus, M., Byrne, D., Bozso, P., Wetmore, D.Z., Mariani, T.J., Kohane, I.S., and Szallasi, Z. 2004. Sequence-matched probes produce increased cross-platform consistency and more reproducible biological results in microarray-based gene expression measurements. Nucleic Acids Res. 32:e74. Miller, R.M. and Federoff, H.J. 2006. Microarrays in Parkinson’s disease: A systematic approach. NeuroRx 3:319-326.

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Mitalipova, M., Calhoun, J., Shin, S., Wininger, D., Schulz, T., Noggle, S., Venable, A., Lyons, I., Robins, A., and Stice, S. 2003. Human embryonic stem cell lines derived from discarded embryos. Stem Cells 21:521-526.

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Miura, T., Luo, Y., Khrebtukova, I., Brandenberger, R., Zhou, D., Thies, R.S., Vasicek, T., Young, H., Lebkowski, J., Carpenter, M.K., and Rao, M.S. 2004. Monitoring early differentiation events in human embryonic stem cells by massively parallel signature sequencing and expressed sequence tag scan. Stem Cells Dev. 13:694-715.

Shamblott, M.J., Axelman, J., Littlefield, J.W., Blumenthal, P.D., Huggins, G.R., Cui, Y., Cheng, L., and Gearhart, J.D. 2001. Human embryonic germ cell derivatives express a broad range of developmentally distinct markers and proliferate extensively in vitro. Proc. Natl. Acad. Sci. U.S.A.98:113-118.

Novoradovskaya, N., Whitfield, M.L., Basehore, L.S., Novoradovsky, A., Pesich, R., Usary, J., Karaca, M., Wong, W.K., Aprelikova, O., Fero, M., Perou, C.M., Botstein, D., and Braman, J. 2004. Universal Reference RNA as a standard for microarray experiments. BMC Genomics 5:20.

Shi, L., Tong, W., Fang, H., Scherf, U., Han, J., Puri, R.K., Frueh, F.W., Goodsaid, F.M., Guo, L., Su, Z., Han, T., Fuscoe, J.C., Xu, Z.A., Patterson, T.A., Hong, H., Xie, Q., Perkins, R.G., Chen, J.J., and Casciano, D.A. 2005. Cross-platform comparability of microarray technology: Intraplatform consistency and appropriate data analysis procedures are essential. BMC Bioinformatics 6:S12.

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Gallagher, K., Ge, W., Guo, L., Guo, X., Hager, J., Haje, P.K., Han, J., Han, T., Harbottle, H.C., Harris, S.C., Hatchwell, E., Hauser, C.A., Hester, S., Hong, H., Hurban, P., Jackson, S.A., Ji, H., Knight, C.R., Kuo, W.P., LeClerc, J.E., Levy, S., Li, Q.Z., Liu, C., Liu, Y., Lombardi, M.J., Ma, Y., Magnuson, S.R., Maqsodi, B., McDaniel, T., Mei, N., Myklebost, O., Ning, B., Novoradovskaya, N., Orr, M.S., Osborn, T.W., Papallo, A., Patterson, T.A., Perkins, R.G., Peters, E.H., Peterson, R., Philips, K.L., Pine, P.S., Pusztai, L., Qian, F., Ren, H., Rosen, M., Rosenzweig, B.A., Samaha, R.R., Schena, M., Schroth, G.P., Shchegrova, S., Smith, D.D., Staedtler, F., Su, Z., Sun, H., Szallasi, Z., Tezak, Z., Thierry-Mieg, D., Thompson, K.L., Tikhonova, I., Turpaz, Y., Vallanat, B., Van, C., Walker, S.J., Wang, S.J., Wang, Y., Wolfinger, R., Wong, A., Wu, J., Xiao, C., Xie, Q., Xu, J., Yang, W., Zhang, L., Zhong, S., Zong, Y., and Slikker, W. 2006. The MicroArray Quality Control (MAQC) project shows inter- and intraplatform reproducibility of gene expression measurements. Nat. Biotechnol. 24:1151-1161. Shin, S. and Rao, M.S. 2006. Large-scale analysis of neural stem cells and progenitor cells. Neurodegener Dis. 3:106-111. Situma, C., Hashimoto, M., and Soper, S.A. 2006. Merging microfluidics with microarray-based bioassays. Biomol. Eng. 23:213-231. Skottman, H., Mikkola, M., Lundin, K., Olsson, C., Stromberg, A.M., Tuuri, T., Otonkoski, T., Hovatta, O., and Lahesmaa, R. 2005. Gene expression signatures of seven individual human embryonic stem cell lines. Stem Cells 23:13431356. Suhr, S.T. and Gage, F.H. 1993. Gene therapy for neurologic disease. Arch. Neurol. 50:12521268. Suhr, S.T. and Gage, F.H. 1999. Gene therapy in the central nervous system: The use of recombinant retroviruses. Arch. Neurol. 56:287-292. Takahashi, K. and Yamanaka, S. 2006. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126:663-676. Tatematsu, M., Tsukamoto, T., and Inada, K. 2003. Stem cells and gastric cancer: Role of gastric and intestinal mixed intestinal metaplasia. Cancer Sci. 94:135-141. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

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Microarray Analysis of Stem Cell Gene Expression

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Current Protocols in Stem Cell Biology

Phenotypic Analysis of Human Embryonic Stem Cells

UNIT 1B.3

Mark Ungrin,1 Michael O’Connor,2,3 Connie Eaves,2 and Peter W. Zandstra1 1

IBBME, University of Toronto, Toronto, Canada Terry Fox Laboratory, BC Cancer Agency, Vancouver, British Columbia 3 StemCell Technologies, Inc, Vancouver, British Columbia 2

ABSTRACT Human embryonic stem cells (hESCs) are an important tool for the study of developmental biology and may one day serve as a source of cells for regenerative medicine. As no definitive assay for hESC pluripotency is available, surrogate assays that measure markers or properties that have been correlated with hESC developmental potential are used to measure the effects of test conditions on their propagation and differentiation. This unit presents a range of protocols, including visual inspection, flow cytometry, immunofluorescence, quantitative real-time reverse-transcriptase PCR, and a colonyforming assay, as tools to measure the undifferentiated hESC state. The authors discuss the advantages and limitations of the various protocols, and present expected results and discuss potential problems. The development of quantitative assays of hESC developmental potential are critical for our understanding of hESC biology. Curr. Protoc. Stem C 2007 by John Wiley & Sons, Inc. Cell Biol. 2:1B.3.1-1B.3.25.
Keywords: hESC r marker r Oct4 r SSEA-3 r SSEA-4 r Tra-1-60 r microscopy r flow cytometry r quantitative real-time PCR

INTRODUCTION Effective techniques for the controlled propagation and differentiation of human embryonic stem cells (hESCs) are prerequisites for the use of these cells for many clinical, industrial, and research applications. Presently however, our ability to develop such methods is limited by a poor understanding of the biology of hESCs. In fact, what are routinely referred to as cultures of “undifferentiated” hESCs comprise complex mixtures of differentiated and undifferentiated cell types. This is due to the variable levels of spontaneous differentiation that result from the widespread use of incompletely defined and/or controlled hESC maintenance conditions. Improvement of conditions requires robust assays to both assess the status of the cells present and provide quantitative information about the relative proportion of undifferentiated and differentiated cells within a given culture. As hESC maintenance cultures typically require daily medium changes, visual examination to look for changes in cell and colony morphology (Basic Protocol 1) is a common approach for monitoring hESC cultures. Although nonquantitative, this method can provide a sensitive and useful low-cost method for the early detection of changes in undifferentiated cell content without disturbing the cells. Expression of a number of more specific phenotypic markers have now been found to correlate with the undifferentiated hESC state, and detection of these lend themselves to more quantitative data acquisition. These phenotypic markers range from cell surface antigens like the stage-specific embryonic antigens (SSEA3 and 4) and the tumor rejection antigens (Tra-1-60 and Tra-1-81) to alkaline phosphatase (Thomson et al., 1998) to various transcription factors including Pou5F1/Oct3/4 (POU domain, class 5, transcription factor 1/Octamer binding transcription factor 3/4), Nanog, teratocarcinoma-derived growth factor-1 (TDGF-1), and CD90

Current Protocols in Stem Cell Biology 1B.3.1-1B.3.25 Published online August 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b03s2 C 2007 John Wiley & Sons, Inc. Copyright


Embryonic and Extraembryonic Stem Cells

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Phenotypic Analysis of hESCs

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Table 1B.3.1 Comparison of Pluripotency Monitoring Techniques

Time required

Visual hESC colony-forming observation cell assay

Flow cytometry for surface antigens

Automated Flow cytometry for Immunofluorescence Quantitative real-time immunofluorescence intracellular antigens microscopy RT-PCR microscopy

Minutes

2 hr

3 hr

1 daya

3 daysa

5 hr

b

b

1 hr

Current Protocols in Stem Cell Biology

Can recover live cells?

Yes

Yes

Yes

No

No

No

No

SSEA, Tra antigen detection?

No

Possible

Yes

Yes

Yes

Yes

No

Oct4 detection?

No

Possible

No

Yes

Yes

Yes

Yes

Alkaline phosphatase detection?

No

Yes

Possiblec

Possiblec

Possible

Possible

Possible

Cell-by-cell data?

No

Yes

Yes

Yes

Yes

Yes

No

Subcellular No localization data?

No

No

No

Yes

Yes

No continued

Current Protocols in Stem Cell Biology

Table 1B.3.1 Comparison of Pluripotency Monitoring Techniques, continued

Visual hESC colony-forming observation cell assay

Flow cytometry for surface antigens

Automated Flow cytometry for Immunofluorescence Quantitative real-time immunofluorescence intracellular antigens microscopy RT-PCR microscopy

Multiple conditions in parallel?

++

++

++

++

+

+++

+++

Sample lifetime

Immediate

Weeks

Immediate

Up to 1 week

Up to 1 week

Up to 1 week

Months to years

Cell/colony positional data?

Yes

Yes

No

No

Yes

Yes

No

Typical use

Routine daily monitoring of cultures

Routine weekly monitoring of cultures; quantitative assessment of pluripotent cells during hESC differentiation

Weekly or monthly monitoring of cultures, response to different culture techniques

Weekly or monthly monitoring of cultures, response to different culture techniques

Marker analysis for a Basic marker Differentiation limited number of analysis for a large assessment, storage for conditions, number of conditions retrospective analysis high-resolution subcellular localization

a Immunofluorescence microscopy protocols also require advance cultivation of cells on appropriate coverslips or multi-well plates. b Protocols could potentially be modified for live-cell imaging of surface antigens. c While the detection of alkaline phosphatase activity by flow cytometric means has been previously reported (He and Landau, 1995), the authors do not have direct experience with this approach. Note also that various antibodies to alkaline phosphatase are also available.

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Supplement 2

(see Henderson et al., 2002; Cai et al., 2006; Trounson, 2006; and the International Stem Cell Initiative; ISCI,http://www.stemcellforum.org/). Techniques for quantitative phenotypic analysis include flow cytometry (FC; Basic Protocol 2 and Alternate Protocol 1), quantitative real-time polymerase chain reaction (Q-RT-PCR; Basic Protocol 4), colony-forming assays (Basic Protocol 5), and immunofluorescence (IF; Basic Protocol 3 and Alternate Protocol 2). Q-RT-PCR and IF-based techniques are somewhat more labor-intensive, but can provide more data in return, with some protocols being capable of assessing tens or a few hundreds of conditions in a single experiment. Q-RT-PCR is particularly well suited to the analysis of large numbers of markers from a single sample, while IF can provide not only subcellular localization data, but is also capable of placing the results from single-cell data in the context of spatial information about location within a colony, local cell density, density of specific subtypes, etc. Currently, no single molecular marker on its own seems capable of defining the pluripotent state of hESCs, and there is presently no “assay” that can measure hESC pluripotency at a single-cell level. The closest strategy is to infer hESC pluripotency from the production of multiple germ line descendants in teratomas generated in vivo from every innoculus (Thomson et al., 1998; Reubinoff et al., 2000) or in vitro from analyses of differentiating aggregates derived from large numbers of input hESCs (e.g., Chadwick et al., 2003; Segev et al., 2005; Ji et al., 2006; Xu et al., 2006; Toh et al., 2007). An alternative functional approach to quantify undifferentiated hESCs is to assess their ability, following complete dissociation, to generate colonies of adherent alkaline phosphatase– positive progeny in vitro. The concern with this latter method is that the frequency of the colony-forming cells (CFCs) detected rarely exceeds 1% (Amit et al., 2000; Forsyth et al., 2006). This finding thus raises the possibility that the CFC assay underestimates the number of undifferentiated hESCs that might grow under nondisaggregated conditions.

STRATEGIC PLANNING In this unit the authors discuss methods for the detection of undifferentiated hESCs that involve a range of time requirements and utilities. Table 1B.3.1 summarizes the protocols presented, together with many of the advantages and disadvantages associated with each. Due to the tendency of hESCs to undergo some spontaneous differentiation in standard maintenance cultures, significant planning and routine monitoring is required, substantially more so than is the case with the majority of other tissue culture systems. Moreover, lot-to-lot variability in undefined culture components—e.g., extracellular matrix components or mouse embryonic fibroblast (MEF) feeder populations—can result in sudden, unexpected changes in the distribution of cell types present. Uniform and reproducible passaging of hESCs can thus be difficult to achieve, even for those with experience. As a result, it is often necessary to modify experimental plans to suit the availability of cells, and the use of various monitoring endpoints to define stopping points can often be helpful. The option to prepare RNA and then store it for extensive periods of time prior to Q-RT-PCR analysis is particularly useful in this context, as samples can be routinely prepared as they become available, and then analyzed retrospectively for specific markers chosen at a later time. BASIC PROTOCOL 1

Phenotypic Analysis of hESCs

VISUAL OBSERVATION OF hESC CULTURES Despite limitations in terms of quantification and sensitivity, one of the most common means of assessing the status of hESC cultures remains visual observation (both direct and light-microscope) of unstained cultures by a trained individual. Regions of high density with evidence of three-dimensional cell layering, representing clumps of insufficiently dissociated or dispersed hESCs during passaging, can often be observed with the naked

1B.3.4 Supplement 2

Current Protocols in Stem Cell Biology

Figure 1B.3.1 Visual inspection of hESC culture. Images of undifferentiated (A, C, E) and partially differentiated (B, D, F) colonies of hESC were captured at 4× (A,B), 10× (C,D) and 20× (E,F) magnification. Scale bars represent 200 µm. The tightly packed cells and sharp colony boundary characteristic of undifferentiated hESC colonies are clearly apparent, particularly at higher magnification (C,E). Large, flattened cells (D,F) are characteristically observed where differentiation is occurring (red arrows)–often present in small areas in all cultures, but spreading to engulf the entire culture surface under conditions which do not support pluripotency. Quality assessment can be used for selective passaging. A growth area defined as “high quality” (G) shows an even appearance, many circular cells at high magnification, and at high density appears evenly multilayered and approximately circular in shape with a well defined edge. It contains a tightly-packed arrangement of cuboidal-like cells that have high nucleus to cytoplasm ratio. Growth areas are defined as differentiated (H,I) when they exhibit clearings (white arrow), uneven thickness in the Z axis, or an uneven rim around colony (black arrow).

eye. Similarly, the size and confluence of hESC growth areas are also visually observable and can be used as a guide for assessing when to passage. Under standard brightfield or phase microscopy growth areas of undifferentiated hESCs have a distinctive morphology (Fig. 1B.3.1A, C, and E) that can be discriminated from the differentiated cells they generate (Fig. 1B.3.1B, D, and F). Under commonly employed “nonselective” passaging regimens, these differences reflect the overall culture quality and differentiation status of the hESCs present, and can give an early indication of problems with culture technique. The morphological differences that distinguish undifferentiated and differentiating hESCs can also be exploited in “selective” passaging techniques where only growth areas containing undifferentiated hESCs (Fig. 1B.3.1G) are collected (Schatten et al., 2005) and the remaining “undesired” growth areas (Fig. 1B.3.1H and I) are discarded.

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Supplement 2

BASIC PROTOCOL 2

FLOW CYTOMETRIC MEASUREMENT OF hESC SURFACE ANTIGENS A number of cell surface antigens have been shown to correlate with the undifferentiated hESC state (see Henderson et al., 2002; Trounson 2006 and references therein as well as the International Stem Cell Initiative–http://www.stemcellforum.org.uk/–for more detailed discussion). Examples of markers that have been extensively used and that have been found to correlate with undifferentiated hESC status include the glycolipid antigens SSEA-3 and SSEA-4 and the protein antigen CD90. The availability of antibodies that recognize these pluripotency related antigens provides a robust and straightforward means of characterizing expression of these markers within a given hESC population by flow cytometry (FC). Note that as the primary antibodies for the Tra-1-60, Tra-1-81, SSEA-1, SSEA-3, and SSEA-4 antigens reflect a range of isotypes and host species, and directly conjugated antibodies for some of these antigens are becoming available, it is possible and desirable to perform assays for multiple antigens in a single experiment (this analysis is illustrative as often populations of cells expressing one pluripotency-associated marker, but not another, are identified). As multiple fluorophores are employed, it is necessary to establish the correct detector compensation settings to avoid cross-talk between channels, a process that requires separate controls stained with each fluorophore individually. The authors routinely combine Tra-1-60, SSEA-4, and 7-AAD (for cell viability) staining using the protocol presented below, but many other combinations are also possible. Note also that as cell fixation and permeabilization are not required for detection of these particular antigens, it is possible to sort and recover viable cells after the staining is finished; e.g., Stewart et al. (2006). For more information on detector compensation settings, consult the manual for the flow cytometer control software. In order to validate antibodies and technique, it is useful to employ positive-control cell lines such as the 2102Ep human teratocarcinoma line (Andrews et al., 1982; Josephson et al., 2007), which are more easily maintained as undifferentiated cultures than hESCs. FC results from this line are shown in Figure 1B.3.2, panels B and C. FC results typically obtained from a population of undifferentiated hESCs are shown in Figure 1B.3.2, panels D, E, and G.

Materials Single-cell suspensions of hESCs (Support Protocol) Hank’s buffered saline solution with 2% (v/v) FBS (HF) Appropriate primary and secondary antibodies 7-aminoactinomycin-D (7-AAD), optional DNase, optional 15-ml microcentrifuge tubes 40-µm cell strainer Flow cytometer sample tubes (e.g., BD Falcon 352058) Flow cytometer Additional reagents and equipment for preparing a single-cell suspension of hESCs (Support Protocol) Measure cell surface antigens 1. Prepare a single-cell suspension of hESCs (Support Protocol). 2. Deposit 2 × 105 cells into each 1.5-ml microcentrifuge tube. The authors have observed substantial differences in cell recovery from different brands of microcentrifuge tubes. They currently employ 1.5-ml Microtubes (DiaMed Lab Supplies, no. SPE150-N). Phenotypic Analysis of hESCs

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Current Protocols in Stem Cell Biology

Figure 1B.3.2 Flow cytometric detection of pluripotency markers. (A-C) 2102Ep teratocarcinoma line. Bracketed numbers represent percent positive in control and sample respectively, using the marker region drawn on each histogram. Each panel shows a histogram of flow cytometry results, paired with a control histogram from a sample prepared without primary antibody. Results for Oct4 (9.9%:53.2%) are shown in panel (A), while panel (B) shows SSEA-4 staining (6.2%:98.1%), and Tra-1-60 (10.1%:56.8%) results are shown in panel (C). (D-G) hESC. Panels (D) and (E) show dot plots of the negative control (no primary antibodies) and combined SSEA-4 (horizontal axis) and Tra-1-60 (vertical axis) flow cytometry results. The percentage of cells in each quadrant is shown on the figure. Note that some cells have lost Tra-1-60 expression but remain SSEA-4 positive. Panels (F) and (G) show Oct4 (4.7%:80.5%) and SSEA-3 (3.2%:85.0%) histogram data, respectively (each with its negative control for comparison).

3. Resuspend in 100 µl of HF. Centrifuge 3 min at 800 × g (200 × g if replating after FC is desired), room temperature. DNase can be added at a final concentration of 0.1 mg/ml during this step to help avoid cell aggregation.

Add primary antibody 4. Add appropriate volumes of primary antibody to the test samples and incubate 30 min at 4◦ C. The authors generally use 1 µl of purified antibody solution, or 20 µl of hybridoma supernatant–depending on supplier; however, it is strongly recommended that each new antibody lot be titrated to determine the optimal concentration.

5. Add 1 ml of HF. 6. Centrifuge 3 min at 800 × g (200 × g if replating after FC is desired), room temperature. 7. Remove supernatant but leave 10 to 20 µl to avoid aspirating pellet. 8. Add 100 µl of HF. Current Protocols in Stem Cell Biology

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Add secondary antibody 9. Add appropriate secondary antibody. Generally, 1 µl of the antibody stock is added to the volume of the tube (110 to 120 µl), which will result in a dilution of 1:110 or 1:120. This and subsequent steps should be performed in the dark as much as possible to prevent photobleaching of the fluorophore.

10. Incubate for 30 min at 4◦ C. 11. Wash twice with 1 to 1.5 ml of HF, each time centrifuging for 3 min at 800 × g (200 × g if replating after FC is desired), room temperature. 12. Resuspend cells in 0.5 to 1 ml of HF. 13. Filter the cells through a 40-µm cell strainer into FACS tubes. 14. Optional: Add 1 µl of 1 mg/ml 7-AAD, incubate 15 min at 4◦ C. 15. Examine the cells by flow cytometry according to the equipment manufacturer’s instructions. Use the software provided by the manufacturer or WinMDI (http://facs.scripps.edu/software.html) to analyze the data. ALTERNATE PROTOCOL 1

DETERMINING Oct4 INTRACELLULAR EXPRESSION BY FLOW CYTOMETRY The nuclear transcription factor Pou5F1 (also known as Oct3, Oct4, and Oct3/4; Rosner et al., 1990; Reubinoff et al., 2000) is one of the most widely accepted markers of the undifferentiated ESC state (Boyer et al., 2005) and forms an elaborate auto-regulatory network with the transcription factors SOX2 and Nanog (Chambers et al., 2003; Mitsui et al., 2003). As Pou5F1 is an intracellular (nuclear) protein, cell permeabilization is required for its detection by FC, making this protocol incompatible with the recovery of live cells. Note, however, that this protocol is compatible with the detection by FC of some cell surface markers, thereby allowing staining for Oct4 and surface markers to be combined and correlated. However, for such protocols, it is important to first determine that the cell surface antigen to be detected is insensitive to the fixation/permeabilization treatment or that the antibodies used to label them prior to the fixation/permeabilization treatment are resistant. As with all FC protocols, care must be taken that compensation settings are set appropriately. Note also that due to the permeabilization step, all cells will take up 7-AAD, which in this case may be used to give an indication of DNA content rather than cell viability. Viability assays for fixed cells have also been described (Riedy et al., 1991). Oct4 FC results from the 2101Ep control cells are shown in Figure 1B.3.2, panel A, while results typically obtained from a population of undifferentiated hESCs are shown in Figure 1B.3.2F.

Materials

Phenotypic Analysis of hESCs

Single-cell suspension of hESCs (Support Protocol) Hank’s buffered-saline solution with 2% (v/v) FBS (HF) IntraPrep Permeabilization Kit (Beckman Coulter) containing: Reagent 1 Reagent 2 Primary antibody: Mouse anti–mouse Oct3/4 antibody (IgG1 isotype; BD Biosciences, cat. no. 611202; http://www.bdbiosciences.com) Secondary antibody: Goat anti–mouse IgG (Fc specific)-FITC conjugate (Sigma-Aldrich, cat. no. F-2772; http://www.sigmaaldrich.com/) DNase, optional

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1.5-ml microcentrifuge tubes (DiaMed Lab Supplies, no. SPE150-N) Vortex 40-µm cell strainer FC sample tubes (e.g., BD Falcon 352058) Flow cytometer Additional reagents and equipment for preparing a single-cell suspension of hESCs (Support Protocol) Prepare cells 1. Prepare a single-cell suspension of hESCs (Support Protocol). DNase can be added at a final concentration of 0.1 mg/ml during this step to help avoid cell aggregation.

2. Load 106 cells into each microcentrifuge tube. Note that this protocol can be completed with fewer cells per sample; however, it is important to maintain consistency between samples and controls within a given experiment. The authors have observed substantial differences in cell recovery from different brands of microcentrifuge tubes. They currently employ 1.5-ml microtubes from DiaMed Lab Supplies (cat. no. SPE150-N).

3. Centrifuge 3 min at 800 × g, room temperature. 4. Remove supernatant, but leave 10 to 20 µl to avoid aspirating pellet. 5. Resuspend in 100 µl of HF.

Fix cells 6. Add 100 µl reagent 1 from the IntraPrep Permeabilization kit. CAUTION: Formaldehyde is considered a toxin and possible carcinogen and appropriate precautions should be taken.

7. Vortex briefly to mix. 8. Incubate 15 min at room temperature. Alternatively, at this stage the sample may be left up to several days at 4◦ C, although signal over background may be slightly reduced.

9. Add 1 ml of HF. 10. Centrifuge 3 min at 800 × g, room temperature. 11. Remove supernatant, but leave 10 to 20 µl to avoid aspirating pellet. 12. Add 100 µl of reagent 2 from the IntraPrep Permeabilization kit.

Add primary antibody 13. Add appropriate volume of primary antibody (1 µl if using the anti-Oct4 antibody listed above). If no dilution is specified for the antibody, use 1 µl of purified antibody (a 1:100 dilution) as a starting point.

14. Incubate at room temperature for 20 min. 15. Add 1 ml of HF. 16. Centrifuge 3 min at 800 × g, room temperature. 17. Remove supernatant, but leave 10 to 20 µl to avoid aspirating pellet. 18. Add 100 µl HF.

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Add secondary antibody 19. Add appropriate volume secondary antibody (2 µl if using the anti–mouse IgG antibody listed above). If no dilution is specified for the antibody, use 1 µl of purified antibody (a 1:100 dilution) as a starting point. This and subsequent steps should be performed in the dark as much as possible to prevent photobleaching of the fluorophore.

20. Incubate for 20 min at room temperature. 21. Add 1 ml of HF. 22. Centrifuge 3 min at 800 × g, room temperature. 23. Remove supernatant, but leave 10 to 20 µl to avoid aspirating pellet. 24. Resuspend in 0.5 to 1 ml of HF. 25. Pass through a 40-µm cell strainer into flow cytometer sample tubes. 26. Examine the cells by flow cytometry according to the equipment manufacturer’s instructions. Use the software provided by the manufacturer or WinMDI (http://facs.scripps.edu/software.html) to analyze the data. SUPPORT PROTOCOL

PREPARATION OF A SINGLE-CELL SUSPENSION OF VIABLE hESCS Single-cell suspensions of hESCs are a prerequisite for many of the assay protocols described in this unit.

Materials Cultures of hESCs in 6-well plates (grown to ∼80% or as desired for a specific experiment) TrypLE Express (Invitrogen, no. 12604-013) PBS (Invitrogen) 37◦ C incubator 15-ml conical centrifuge tube Hemacytometer Additional reagents and equipment for counting cells (Phelan, 2006) 1. For hESCs cultured in a 6-well plate, add 1 ml TrypLE per well (scale volume accordingly for other culture surfaces). Trypsin may be substituted for the TrypLE Express, however in this case at step 3, substitute 4 ml of medium containing serum or trypsin inhibitors (depending on subsequent application) for PBS to prevent damage to the cells. Alternatively, colonies in suspension (e.g., after collagenase passaging) may be resuspended in TrypLE, such that only part of a well is required and the remainder may be employed for other purposes.

2. Incubate for 10 min at 37◦ C. 3. Add 4 ml PBS. 4. Triturate to wash cells off culture surface and break up clumps. 5. Transfer to 15-ml conical centrifuge tube. 6. Remove an aliquot to count cells with hemacytometer (Phelan, 2006). Phenotypic Analysis of hESCs

1B.3.10 Supplement 2

7. Centrifuge 5 min at 200 × g, room temperature. 8. Resuspend in PBS to desired density. Current Protocols in Stem Cell Biology

IMMUNOFLUORESCENT STAINING OF FIXED hESCS ON COVERSLIPS Immunofluorescent staining (IF) is a powerful technique for the analysis of cell type as well as spatial location and relationships. This technique is appropriate when a limited number of samples are to be visualized in detail. Cells are fixed and permeabilized in situ, followed by sequential antibody probing as per the FC protocols described above. Importantly, information on the subcellular localization of proteins that is inaccessible using flow cytometric approaches may be obtained using IF. Primary antibodies may be combined as noted above for FC, with appropriate choices of fluorescently labeled secondary antibodies. Additional information may be obtained using any of a large number of fluorescent probes (see the Molecular Probes Handbook at http://probes.invitrogen.com/handbook/ for further information), being limited only by the excitation and detection limits of available microscopes. Note that standard microscope optics are generally calibrated for a no. 1 1/2 coverslip (see http://www.microscopyu.com/articles/formulas/formulascoverslipcorrection.html for discussion). It is possible to seed adherent cells on a microscope slide for imaging, however it is preferable to culture cells directly on the coverslip for optimal performance, particularly for high-magnification applications.

BASIC PROTOCOL 3

Materials Poly-lysine (L or D), optional Phosphate-buffered saline (PBS; Invitrogen) Desired substrate (e.g., Matrigel, mouse embryo fibroblasts) hESCs 3.7% (w/v) formaldehyde in PBS Methanol (chilled to −20◦ C) Appropriate primary and fluorescently labeled secondary antibodies 3% (w/v) BSA in PBS Mounting medium/antifade Cosmetic nail polish Coverslips Glass petri dish Filter paper Beakers Microscope slides Prepare coverslips 1. Optional: Precoat coverslips with poly-lysine to increase adhesion: a. Make filter-sterilized stock of 1 mg/ml poly-lysine in PBS (can be stored as frozen aliquots). b. Dilute to 10 µg/ml in sterile PBS to obtain working solution. c. Immerse coverslips in working solution for 30 min at room temperature. d. Aspirate medium. e. Air dry coverslips and store for months, or use immediately. 2. Prepare coverslips with desired substrate (e.g., Matrigel, mouse embryo fibroblasts, etc.). 3. Seed hESCs at a density appropriate to the desired experiment. 4. Allow cells to grow until desired time point is reached. Keep in mind that if cells have not had time to completely adhere to the substrate, they may detach during the staining protocol.

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Fix cells 5. Aspirate the medium. 6. Add 3.7% formaldehyde in PBS. CAUTION: Formaldehyde is considered a toxin and possible carcinogen, appropriate precautions should be taken.

7. Incubate 5 min at room temperature. 8. Wash three times with PBS (5 min each wash). For extracellular staining: only fix the cells, do not permeabilize (skip steps 9 to 11).

9. Add methanol (−20◦ C). 10. Incubate for 5 min at −20◦ C. 11. Wash three times with PBS (5 min each wash). At this point, samples can be stored in PBS for up to 1 week.

12. Transfer coverslips into a glass petri dish lined with filter paper moistened with distilled water. The filter paper is moistened to prevent the coverslips from drying out during the incubation step, however if it is actually wet, formation of a water drop at the edge of a coverslip can draw off the antibody solution.

Add primary antibody 13. Add primary antibody diluted to 20 µl in 3% BSA in PBS to each coverslip. Surface tension will hold the liquid on the coverslip.

14. Incubate 45 min at 37◦ C. 15. Wash coverslips in PBS by dipping sequentially in three beakers. 16. Transfer onto a moistened dish as above.

Add secondary antibody 17. Add secondary antibody diluted to 20 µl in PBS to each coverslip. 18. Incubate 30 to 45 min at 37◦ C. 19. Wash in PBS sequentially (as above). 20. Mount in mounting medium/antifade by placing a drop on a slide and gently inverting the coverslip cell side down into the medium, while avoiding air bubbles. 21. Let dry for 30 min or more (see instructions for specific brand of mounting medium/antifade). 22. Seal the edges of the coverslip to the slide with nail polish. 23. Examine the cells using a fluorescence microscope equipped with the appropriate optics and filters. ALTERNATE PROTOCOL 2

Phenotypic Analysis of hESCs

IMMUNOFLUORESCENCE STAINING FOR HIGH-CONTENT SCREENING High-content screening employs automated microscopy to monitor marker expression patterns at the single-cell level under a larger number of conditions than might otherwise be possible. It uses sample preparation techniques that are similar to those described above for individual samples on coverslips. Results obtained following this protocol to probe for Tra-1-60 and Oct4 are shown in Figure 1B.3.3.

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Current Protocols in Stem Cell Biology

Figure 1B.3.3 Detection of pluripotency markers by immunofluorescence. Colonies of hESCs were probed for Tra-1-60 (red, A) or Oct4 (green, B) and imaged in a Cellomics ArrayScan VTI automated microscope. Scale bars represent 200 µm. In both cases, Hoechst 33342 dye was also added to label nuclei (blue). Note the clearly defined nuclear staining of Oct4, which makes this marker an ideal choice for automated image segmentation and analysis.

Materials hESCs 3.7% (w/v) formaldehyde in PBS Phosphate-buffered saline (PBS; Invitrogen) Methanol (chilled to −20◦ C) 10% (v/v) FBS in PBS Appropriate primary and secondary antibodies Hoechst 33342 (Sigma-Aldrich, cat. no. B2261), frozen stock at 1 mg/ml Multi-well tissue culture plates designed for use in microscopy (Perkin Elmer, Packard View Plate, cat. no. 6005182; Fisher Scientific, Corning 96-well, cat. no. 07-200-729) Current Protocols in Stem Cell Biology

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Multichannel pipet Disposable reagent reservoirs Sterile trough 200-µl pipet Aluminum foil Cellomics ArrayScan VTI (Cellomics, http://www.cellomics.com, or equivalent automated microscopy platform); see also CellProfiler, a freely available open-source automated image analysis package, http://www.cellprofiler.org/ (Carpenter et al., 2006) Plate cells 1. Prepare culture surface according to desired culture conditions, and passage hESC cells normally onto the 96-well plate. Note that the surface area of one well in these plates is 0.3 cm2 as compared to 9.6 cm2 in the well of a 6-well plate. Thus, 1 well of cells from a 6-well plate passaged to one full 96-well plate involves a 1:3 split ratio. The Corning thin-bottom plate permits somewhat higher sensitivity of detection, however the glass is prone to flexing, which can displace the focal plane of neighboring fields and necessitate more frequent refocusing during imaging. This can be avoided if the user is careful not to allow pipet tips to contact the bottom of the wells.

2. Culture the cells under the desired conditions until ready to image. If overgrowth occurs resulting in cells growing on top of one another, this complicates automated image analysis. For timed exposures, start times should be staggered such that all samples are ready for fixation simultaneously, as fixation of a portion of the wells in the plate has detrimental effects on the remaining live cells.

Fix cells 3. Gently remove the medium using a multichannel pipet. Alternatively, to remove medium quickly at a defined time point, the plate may be inverted on top of a sterile disposable towel and tapped twice.

4. Add 200 µl of a 3.7% formaldehyde solution in water to each well. CAUTION: Formaldehyde is considered a toxin and possible carcinogen, appropriate precautions should be taken.

5. Incubate the plate for 10 min at 37◦ C. 6. Remove fixing reagent using the multichannel pipet, and set aside the tips used to do so. These tips should be considered contaminated with formaldehyde, and disposed of accordingly.

7. Wash three times with PBS (200 µl/well, each wash). The tips and supernatant from the first wash should also be considered contaminated with formaldehyde, and disposed of accordingly.

8. Remove final wash solution using multichannel pipet.

Permeabilize cells 9. Add 200 µl of 100 % methanol to each well. Set a timer at the beginning of methanol addition to 3 min, so that no well is exposed to methanol for longer than 3 min to avoid complete cell lysis. 10. Wash three times with PBS (200 µl/well, each wash). Phenotypic Analysis of hESCs

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Block nonspecific binding sites 11. Remove last wash using a multichannel pipet. 12. Add 200 µl/well 10% FBS in PBS. 13. Incubate overnight at 4◦ C overnight (up to one week).

Add primary antibodies 14. Combine primary antibodies in a master mix in 10% FBS in PBS–make 40 µl per well. For Oct4 detection, the authors use mouse anti–mouse Oct3/4 antibody (IgG1 isotype; BD Biosciences, cat. no. 611202). Antibody dilutions vary; consult the antibody product sheet for recommendations–if no information is available, a 1:200 dilution of concentrated antibody stock may be assayed initially.

15. Remove the blocking solution. 16. Add 38 µl of primary antibody solution to each well individually. Use of the multichannel pipet and reagent reservoir at this stage will result in more wastage of master mix.

17. Incubate overnight at 4◦ C.

Add secondary antibody 18. Combine secondary antibodies and 5 to 10 µg per ml Hoechst 33342 in a master mix in 10% FBS in PBS–make 40 µl per well. Fluorophores that are relatively resistant to photobleaching, such as the Alexa series from Invitrogen, are preferable to minimize quantitation error. Antibody dilutions vary; consult the antibody product sheet for recommendations–if no information is available, a 1:200 dilution of concentrated antibody stock may be assayed initially. Operate under reduced lighting to avoid photobleaching of the secondary antibody.

19. Remove primary antibody solution using multichannel pipet. 20. Wash the plate three times with PBS (200 µl/well each wash), using a multichannel pipet and sterile trough. 21. Add 38 µl of secondary antibody solution to each well individually via a 200-µl pipet (use of the multichannel pipet and reagent reservoir at this stage will require more wastage of master mix). 22. Wrap the plate in aluminum foil, label top of plate, and incubate 1 to 2 hr at room temperature. 23. Under low light, wash the plate three times with PBS (200 µl/well each wash). 24. Add 200 µl/well PBS. 25. Rewrap the plate in aluminum foil and store at 4◦ C until ready to scan. 26. Read the plate within 1 week.

QUANTITATIVE REAL-TIME POLYMERASE CHAIN REACTION (Q-RT-PCR)

BASIC PROTOCOL 4

Numerous manufacturers and systems are available for RNA purification, cDNA generation, primer design, and Q-RT-PCR; the choice of system will be determined by availability and laboratory preference. An in-depth review of Q-RT-PCR analysis techniques is beyond the scope of this unit, but available in many other publications (e.g.,

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1B.3.15

Wong and Medrano, 2005; Lutfalla and Uze, 2006). Here the authors briefly describe a set of protocols used in their laboratories to obtain useful data from hESCs. The general procedure involves first extracting the RNA which is then employed as a template to generate cDNA via a reverse-transcriptase reaction. This cDNA is then subjected to PCR amplification, with real-time monitoring of product levels to give a quantitative measure of the initial concentration of the cDNA (and hence RNA) for the sequence amplified by the primers chosen. For detection of undifferentiated cells the authors routinely use primers against Pou5F1 (Oct4), Nanog, TDGF1, DNM3TB, and UTF1 (see also the International Stem Cell Initiative–http://www.stemcellforum.org.uk/). Experience with multiple differentiation methods shows that these genes are consistently down-regulated upon hESC differentiation. Similarly, for detection of lineage-specific differentiation, the authors have found that the putative lineage markers AFP (endoderm), Hand1 and MSx1 (mesoderm), and Msi1 (ectoderm) are often upregulated upon hESC differentiation. To generate Q-RTPCR primers for specific human genes of interest, the authors use the UCSC genome browser (http://genome.ucsc.edu/) to obtain genomic sequence information showing exon locations, then use Primer3 (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3 www.cgi) for primer design. To determine primer specificity the authors use the NCBI BLAST tool (search for short, nearly exact matches) to compare the putative primer sequences against both the human and mouse genome. New primers are designed if the putative primers show significant homology to genes other than the gene of interest.

Materials hESC cultures Trizol reagent DEPC-treated water 250 ng/µl random hexamers 5× FS buffer RNase inhibitor 0.1 M DTT dNTP mix Superscript II reverse transcriptase cDNA samples Primer pairs (forward and reverse primers can be combined into one working stock with each primer at 20 pmoles/µl; see Table 1B.3.2) SYBR Green Master Mix Nuclease-free tubes GeneAmp PCR System 9700 (or equivalent) ABI 96-well optical reaction plate and adhesive cover Applied Biosystems 7500 Real Time PCR System (or equivalent) Additional reagents and equipment for measuring RNA concentration (Gallagher and Desjardins, 2006) Synthesize cDNA NOTE: Keep all reagents on ice during set up. 1. Harvest hESC sample(s) using Trizol as per the manufacturer’s instructions, then store at −80◦ C. The hESC samples should reach a confluency of ∼70% to 80% (∼106 cells/ml; it is possible to reduce this number: see directions for Trizol as provided by the manufacturer). Phenotypic Analysis of hESCs

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Table 1B.3.2 Primers Used for the Most Commonly Studied Gene Productsa

Gene symbol

Accession number

Forward primer

Reverse primer

GAPDH

NM 002046

CCCATCACCATCTTCCAGGAG

CTTCTCCATGGTGGTGAAGACG

Pou5F1

NM 002701

GTGGAGGAAGCTGACAACAA

CTCCAGGTTGCCTCTCACTC

Nanog

NM 024865

AACTGGCCGAAGAATAGCAA

CATCCCTGGTGGTAGGAAGA

TDGF1

NM 003212

CTGCTTTCCTCAGGCATTTC

TGCAGACGGTGGTAGTTCTG

UTF1

NM 003577

CGCCGCTACAAGTTCCTTA

ATGAGCTTCCGGATCTGCT

AFP

NM 001134

GTAGCGCTGCAAACAATGAA

TCTGCAATGACAGCCTCAAG

Hand1

NM 004821

AACTCAAGAAGGCGGATGG

CGGTGCGTCCTTTAATCCT

Msx1

BC067353

CGAGAAGCCCGAGAGGAC

GGCTTACGGTTCGTCTTGTG

Msi1

NM 002442

CTTTGATTGCCACAGCCTTC

ACTCGTGGTCCTCAGTCAGC

a Due to the presence of pseudogenes for some of these genes, some of the above primer sequences also match pseudogene sequences.

2. Determine RNA concentration (Gallagher and Desjardins, 2006) and purity using your preferred method (the authors employ an Agilent 2100 Bioanalyzer as per the manufacturer’s instructions). 3. To the required number of nuclease-free tubes, add: DEPC-treated water 250 ng/µl random primers purified RNA

Sample tubes 2 µl 1 µl 7 µl

’No RNA control’ tube 9 µl 1 µl –

4. Place tubes containing the primer annealing mix inside GeneAmp PCR System 9700, enter cycle conditions for primer annealing (65◦ C for 10 min, 4◦ C for 15 min, 4◦ C hold), then run annealing program. 5. Make up sufficient RT reaction mix to allow the following volumes per reaction:

4 µl 5× FS buffer 1 µl RNase inhibitor 2 µl 0.1 M DTT 2 µl dNTP 1 µl Superscript II reverse transcriptase (add after all other reagents are added). Vortex to mix, briefly spin down, and then put on ice. 6. Once the primer annealing mix has reached the 4◦ C hold, add 10 µl of the RT-reaction mix to each tube. 7. Place tubes inside GeneAmp PCR System 9700, enter cycle conditions for RT (42◦ C for 60 min, 70◦ C for 10 min, 4◦ C hold), then run RT program. 8. Once the RT reaction has reached 4◦ C, transfer the cDNA samples into appropriately labeled 1.5-ml nuclease-free tubes and store at −20◦ C.

Perform Q-RT-PCR NOTE: Keep all reagents on ice during the set up. 9. Organize plate layout for chosen cDNA samples and primer pairs (including a housekeeping gene for relative quantification; e.g., GAPDH).

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10. Make a master mix of each primer pair in nuclease-free tubes: (n + 1) × 0.5 µl (n + 1) × 4.5 µl (n + 1) × 5.0 µl.

Primers DEPC-water Total

Vortex to mix prior to briefly spinning down. n = the number of reactions to be done per primer pair, including the ‘No RNA control’ from the reverse transcription and a ‘No cDNA control’ for the real-time PCR.

11. Make a master mix of each cDNA sample in nuclease-free tubes using Table 1B.3.3. 12. To an ABI 96-well optical reaction plate add the diluted primer pairs to the appropriate wells. 13. Add SYBR Green Master Mix to the appropriate cDNA-containing tubes. 14. Carefully place an optical adhesive cover onto the 96-well plate and seal, without touching the clear area. 15. Spin the plate briefly at 100 × g to ensure all solutions are at the bottom of each well. 16. Place the 96-well plate inside an Applied Biosystems 7500 Real Time PCR System then enter primer and sample details as prompted, taking care to note the reference gene for relative quantification. 17. Enter Q-RT-PCR conditions (25 µl sample volume), then run program as follows: 50◦ C 95◦ C 95◦ C 60◦ C.

2 min 10 min 15 sec 1 min

40-50 cycles:

18. Once the program has finished running, follow prompts to generate Dissociation Curve. 19. Once the Dissociation Curve program has run, check each well for: a. primer dimer peak around 75◦ C b. single peak around 85◦ C c. peak around 95◦ C indicating genomic DNA contamination. Optional: when using Q-RT-PCR primer pairs for the first time, it is useful to run the Q-RT-PCR reaction products on an agarose gel to confirm that the amplicon is of the expected size. Table 1B.3.3 Master Mix

Phenotypic Analysis of hESCs

cDNA samples

No RNA control

No cDNA control

cDNA

(n + 1) × 0.5 µl

(n + 1) × 0.5 µl

DEPC-water

(n + 1) × 7.0 µl

(n + 1) × 7.0 µl

(n + 1) × 7.5 µl

SYBR Green Master Mixa

(n + 1) × 12.5 µl

(n + 1) × 12.5 µl

(n + 1) × 12.5 µl

Total

(n + 1) × 20.0 µl

(n + 1) × 20.0 µl

(n + 1) × 20.0 µl

—

a Add Sybr Green Master Mix immediately prior to beginning Step 13.

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Current Protocols in Stem Cell Biology

20. If the dissociation curves and/or agarose gel indicate specific product amplification during the Q-RT-PCR reaction, analyze the data following the Relative Quantification Study prompts.

hESC COLONY FORMING CELL ASSAY The colony-forming cell (CFC) assay detects the capacity of single hESCs to form colonies of at least 30 undifferentiated hESC progeny using alkaline phosphatase activity as a surrogate indicator of their ability to execute at least 5 self-renewal divisions.

BASIC PROTOCOL 5

Materials hESCs cultured in 6-well plates TrypLE Express (Invitrogen, cat. no. 12604-013) Phosphate-buffered saline (PBS; Invitrogen) 35-mm dish containing mouse embryonic fibroblasts or Matrigel with MEF-conditioned medium Alkaline phosphatase detection kit (Sigma-Aldrich, cat. no. 86R-1KT). 15-ml conical tubes 40-µm cell strainer Hemacytometer Standard optical microscope Additional reagents and solutions for counting cells using a hemacytometer (Phelan, 2006) CFC assay 1. For hESCs cultured in a 6-well plate, add 1 ml TrypLE per well (scale volume accordingly for other culture surfaces). Trypsin may be substituted for the TrypLE Express, however in this case at step 3, substitute 1 ml of medium containing trypsin inhibitors (not serum) for PBS, to prevent damage to the cells. Alternatively, colonies in suspension (e.g., after collagenase passaging) may be resuspended in TrypLE, such that only part of a well is required and the remainder may be employed for other purposes.

2. Incubate 10 min at 37◦ C. 3. Gently scrape/wash cells off the culture surface then transfer to a 15-ml conical tube. 4. Wash the culture surface with 2 ml of PBS then add this to the 15-ml tube and gently triturate. 5. Filter cells through a 40-µm cell strainer to remove residual cell aggregates. 6. Remove an aliquot to count cells with hemacytometer (Phelan, 2006). 7. Centrifuge 5 min at 200 × g, room temperature. 8. Plate from 104 to 105 cells into a 35-mm dish containing either mouse embryonic fibroblasts (MEF) or Matrigel with MEF-conditioned medium and culture for 7 days.

Fix and stain colonies 9. Fix and stain colonies for alkaline phosphatase using appropriate kit (e.g., Sigma). 10. Count the number of alkaline phosphatase-positive colonies containing >30 cells and determine the hESC CFC frequency within the input cells.

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Supplement 2

COMMENTARY Background Information

Phenotypic Analysis of hESCs

Oct4, also known as Oct3 or Pou5f1, is a transcription factor, and one of the most commonly used markers for undifferentiated ESCs. The Oct4 gene has been cloned and sequenced (Rosner et al., 1990; Reubinoff et al., 2000), and is thus available for detection using both protein and mRNA methods (although the existence of several intron-less pseudogenes necessitates the elimination of contaminating genomic DNA from samples prior to analysis by Q-RT-PCR). There is a substantial body of evidence supporting a role for Oct4, along with two additional transcription factors, Nanog and Sox2 (Boiani and Sch¨oler, 2005; Boyer et al., 2005), in a genetic regulatory loop responsible for the maintenance of ESC pluripotency. While Oct4 is involved in the maintenance of undifferentiated hESC, it has also been observed in other cell types (Takeda et al., 1992; Niwa et al., 2000). The use of Oct4 as a pluripotency marker is thus context-dependent and should be supported by co-expression with other markers such as Nanog (Chambers et al., 2003; Mitsui et al., 2003) and Sox2 (Avilion et al., 2003). Other markers commonly used to measure the undifferentiated hESC state are SSEA-3, SSEA-4, Tra-1-60, and Tra-1-81, (Henderson et al., 2002). Most of these markers were identified empirically as antigens expressed on the surface of human embryonal carcinoma (EC) cells, a transformed cell type with some characteristics of ESC (see Draper et al., 2002; Josephson et al., 2007 and references therein). These markers are nonprotein antigens composed of glycolipids (e.g., SSEA1, SSEA-3, SSEA-4) or keratan sulfates (Tra1-60 and Tra-1-81); the Tra antigens have recently been reported to be associated with the protein podocalyxin (Schopperle and Dewolf, 2006). The biological basis for their regulation and their role in the maintenance of pluripotency is poorly understood. Nevertheless, loss of these markers has been shown to correlate with loss of hESC developmental potential. SSEA-3 expression is particularly interesting as it seems to be lost more rapidly than other commonly used markers, even allowing for the detection of SSEA-3 negative Oct4 positive subpopulations (Enver et al., 2005; Stewart et al., 2006). SSEA-4 is lost more gradually, while the Tra-1-60 and Tra-1-81 antigens exhibit an intermediate behavior (Draper et al., 2002). As these markers are located on the exterior surfaces of the cell membrane, they may be detected without damaging the cells, per-

mitting sorting of live cells for subsequent culture and analysis (Stewart et al., 2006). Many more gene (e.g., TDGF1, FLJ10884, RPC32, NTS, LEFTB, DNMT3B; Enver et al., 2005) and extracellular antigen (TRA-2-49, TRA-254; Andrews et al., 1984; GCTM2, GCTM3; Pera et al., 1988; CD9; Oka et al., 2002; CD90/Thy-1; Draper et al., 2002) markers associated with the hESC state continue to be identified. Ultimately, functional assays which correlate with hESC developmental potential will need to be used to tease apart the biological properties of these markers.

Critical Parameters For visual observation, a consistent microscope set up, including the objective and illumination conditions used are essential to consistently discriminate changes in hESC behavior. If multiple microscopes are to be used, it is worth designating one microscope in particular for visual inspection of hESC cultures. Having one member of the laboratory staff concentrate on developing this expertise, ideally via hands-on training in a facility with prior experience, is also likely to be more effective than trying to train a number of part-time users simultaneously. This “superuser” can then train other personnel, and the necessarily subjective nature of direct visual observation will not be compounded by inconsistencies between individually developed techniques. The authors have observed cases where the same cells, cultured in the same growth medium by different individuals, nevertheless exhibit changes in behavior that may be due to differences in colony size and concomitant perimeter-to-area ratios (M. Ungrin, unpub. observ.). Consequently, in order to obtain reproducible results, it is particularly important to carefully observe and document these parameters. Users will naturally attain increased consistency as habits develop around routine tissue culture operations; however, cells cultured by two experienced individuals may still show differences depending on the specific techniques adopted (i.e., collagenase versus partial trypsinization, degree of trituration, incubation times, density at passage, numbers of cells or growth areas transferred, etc.). Thus every effort should be made to employ cells with similar passage history for any experimental comparisons. For the FC and IF protocols, appropriate selection of antibodies can be a more complex problem than might be expected when multiple

1B.3.20 Supplement 2

Current Protocols in Stem Cell Biology

targets are assayed simultaneously. Fluorophores must be selected based on the availability of appropriate detectors and filters, such that they minimize cross-talk between detection channels. For example, many flow cytometers are not equipped with a laser that has a wavelength shorter than 488 nm, eliminating the use of fluorophores that require excitation in this range. Susceptibility to photobleaching is also an issue, particularly for microscope detection of IF or where intensity quantification is to be attempted. Primary antibodies should differ in species or isotype, both from one another and from the secondary antibodies that will be employed subsequently. For Q-RT-PCR, it is essential that all primer sequences be individually verified before ordering and using them, and there are a number of useful tools available on the internet for this purpose (see Basic Protocol 5). In the authors’ experience, it is not uncommon for published primers to contain one or more sequence errors, and if these are not identified prior to ordering, much time and effort can be wasted. While publicly accessible databases (e.g., Primer Bank, http://pga.mgh.

harvard.edu/primerbank/) are a useful resource, it is advantageous to obtain primer sequences from individuals with whom one has personal contact and can easily approach for troubleshooting assistance if necessary.

Troubleshooting See Tables 1B.3.4, 1B.3.5, 1B.3.6, and 1B.3.7 for information on troubleshooting these protocols.

Anticipated Results Visual observations Visual observations are by their nature more qualitative than the other assays described here; however, the results can, nevertheless, be highly reproducible. The characteristic morphology of a classical hESC colony, with its close-packed cells and sharply defined boundaries (see Fig. 1B.3.1A,C,E) clearly differs from the elongated shapes of the MEF feeder cells, both of which differ from the sheet-like cells that often arise at the periphery of differentiating growth areas (see Fig. 1B.3.1B,D,F). A wide range of other morphologies can also

Table 1B.3.4 Troubleshooting Guide for Flow Cytometry Protocols

Problem

Possible cause

Solution

Cells lost during protocol

Pellet aspirated

Leave a small amount of liquid over the pellet when aspirating

Poor pelleting characteristics of the specific brand of microcentrifuge tubes employed

Test another brand of microcentrifuge tubes

Cells fragmenting

Reduce speed and time to 2 min at 400 × g

Excessive debris in forward-scatter/side-scatter plot

Reduce speed or duration of vortexing in step 7 (Alternate Protocol 1) Excessive background staining

Antibody used at too high a concentration

Decrease concentration of primary and secondary antibodies (alone and in combination)

Weak signal

Fluorophore photobleached

Ensure cells are kept in the dark after addition of fluorescently labeled secondary antibodies

Antibody used at too low a concentration

Increase concentration of primary and secondary antibodies (alone and in combination)

Too many cells

Check accuracy of cell counting

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Table 1B.3.5 Troubleshooting Guide for Immunofluorescence Microscopy

Problem

Possible cause

Solution

Cells lost during protocol

Excessive agitation during wash/pipetting steps

Pipet slowly during antibody addition, transfer slowly into and out of PBS wash step

Cells not given enough time Allow cells to grow on coverslip for at to attach least 24 hr before use Cells do not adhere strongly Precoat coverslips with poly-lysine, enough increase concentration if necessary Uneven staining

Photobleaching of fluorophore

Protect plate from light during procedure Use fluorophores less susceptible to photobleaching

Excessive background staining

No signal

Antibody used at too high a Decrease concentration of primary and concentration secondary antibodies (alone and in combination) Antibody is not specific or not effective for IF

Contact supplier’s technical support line

Antibody used at too low a concentration

Increase concentration of primary and secondary antibodies (alone and in combination)

Consistent image Optics dirty or misaligned artifacts between fields

Consult manual for your microscope

Table 1B.3.6 Troubleshooting Guide for Q-RT-PCR

Problem

Possible cause

Solution

No amplification of positive controls

Poor RNA quality

Check RNA via Agilent chip or gel

Poor cDNA quality

Repeat reverse-transcriptase reaction

Insufficient Q-RT-PCR cycles Perform 40 to 50 cycles Incorrect primers Primer dimmers

Inappropriate primer selection Redesign primers

Incorrect amplicon size

Nonspecific primer binding

be seen, depending on the specific experimental conditions being investigated. It is also important to observe multiple growth areas before drawing conclusions about the differentiation status of a culture, as differentiated and undifferentiated cells can co-exist, and it is the relative balance between the two that is diagnostic. In selective passaging, the overall shape of the growth area as well as its symmetry is also important (see Fig. 1B.3.1G,H,I). Phenotypic Analysis of hESCs

Check primer sequence; redesign primers.

Preparation of single cells The suspension of cells should be free of cell aggregates. This may be difficult to

Redesign primers

achieve with higher-density cultures, where one or more large aggregates may appear. When this occurs, additional trituration, DNase treatment and/or filtration may be necessary. Flow cytometric analysis of hESC cell surface and intracellular antigens Example data from the 2102Ep human teratocarcinoma line (Andrews et al., 1982; Josephson et al., 2007), which expresses some markers of pluripotency and may be employed as a positive control, is shown in Figure 1B.3.2, panels A through C. Data obtained from a

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Current Protocols in Stem Cell Biology

Table 1B.3.7 Troubleshooting Guide for CFC Assay

Problem

Possible cause

Low CFC frequency in undifferentiated hESC cultures

Operator inexperience; hESCs hESCs should be plated left in suspension for too long immediately following generation of single-cell suspension

Faint alkaline phosphatase staining in colonies

Solution

Poor-quality hESC cultures

Passage hESCs selectively, based on morphological criteria for undifferentiated hESCs

Poor-quality MEFs or MEF-conditioned media for CFC assay

Screen Knock-Out Serum Replacer and/or mouse embryonic fibroblasts to find batches suitable for maintaining undifferentiated hESCs

Poor-quality MEFs or MEF-conditioned media

Screen Knock-Out Serum Replacer and/or mouse embryonic fibroblasts to find batches suitable for maintaining undifferentiated hESCs

Over fixation of colonies

Reduce colony fixation time.

Too much time between colony fixation and colony staining

Stain colonies immediately after fixation.

population of hESCs is shown in Figure 1B.3.2 panels D through G. Note from the scatter plot in Figure 1B.3.2E that some cells have lost expression of Tra-1-60 (vertical axis) but still retain expression of SSEA-4. Immunofluorescence analysis of hESCs Typical IF results for hESCs are shown in Figure 1B.3.3. Note the extremely clear Oct4 signal in panel B, making this marker a good choice for automated image analysis applications. Quantitative real-time polymerase chain reaction analysis of hESCs Monitoring the response of hESCs to altered culture conditions or specific differentiation protocols via Q-RT-PCR can provide a great deal of information relating to maintenance of pluripotency-related genes and/or induction of differentiation-related genes. Of the different candidate pluripotency-related genes cited in the literature, the authors have found Oct4, Nanog, TDGF1, and UTF1 to be the most reliably down-regulated using multiple differentiation protocols. As with the CFC protocol, it is important to realize that conditions that promote differentiation of hESCs do not necessarily result in pronounced, immediate changes to cell status and markers.

Colony-forming cell assay Colonies that arise 7 days after plating suspensions of single hESCs are generally compact and stained evenly for alkaline phosphatase. Although variations in colony shape and staining density are observed, the authors have not seen many alkaline phosphatase– negative colonies when cells from either undifferentiated hESC cultures, or from embryoid bodies or retinoic acid-treated hESCs are assayed, thereby rationalizing the utility of this assay for quantifying residual undifferentiated hESCs during differentiation protocols. For karyotypically normal undifferentiated hESCs cultured on mouse embryonic fibroblasts (MEFs) or MEF-conditioned media, CFC frequencies range in the order of 1/100 to 1/1000 cells, depending on the cell line used, operator experience, and quality of culture conditions. Note also the fact that CFCs persist under these conditions for significant periods of time.

Time Considerations Time considerations are a substantial concern in hESC studies. The daily re-feeding required under most culture protocols means maintenance of any significant number of hESC cultures is a significant commitment in and of itself. Moreover, in spite of the best

Embryonic and Extraembryonic Stem Cells

1B.3.23 Current Protocols in Stem Cell Biology

Supplement 2

efforts to maintain consistency, considerable variability is encountered by most groups. Rapid and easily performed methods to assess hESCs are therefore critical. Visual observation clearly meets this need. Live-cell FC analysis of surface markers is also relatively simple and rapid assuming access to a flow cytometer is not limiting. FC of fixed cells can circumvent the latter by allowing samples to be stored for later investigation but then the immediacy of the data is sacrificed. IF techniques require that initiation, maintenance, and fixation of cultures on an appropriate surface occur on a schedule dictated by the cells themselves. Q-RT-PCR techniques take more time to complete, but initial sample collection is rapid, and once the RNA has been prepared, it can be stored at −80◦ C for extended periods of time. Further, as the specific genes targeted for analysis need not be selected until the samples are analyzed, this approach is ideally suited to experiments where the final outcome may not be known and retrospective data collection offers a unique advantage.

Literature Cited Amit, M., Carpenter, M.K., Inokuma, M.S., Chiu, C.P., Harris, C.P., Waknitz, M.A., ItskovitzEldor, J., and Thomson, J.A. 2000. Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227:271278. Andrews, P.W., Goodfellow, P.N., Shevinsky, L.H., Bronson, D.L., and Knowles, B.B. 1982. Cellsurface antigens of a clonal human embryonal carcinoma cell line: Morphological and antigenic differentiation in culture. Int. J. Cancer 29:523-531. Andrews, P.W., Meyer, L.J., Bednarz, K.L., and Harris, H. 1984. Two monoclonal antibodies recognizing determinants on human embryonal carcinoma cells react specifically with the liver isozyme of human alkaline phosphatase. Hybridoma 3:33-39.

Phenotypic Analysis of hESCs

Cai, J., Chen, J., Liu, Y., Miura, T., Luo, Y., Loring, J.F., Freed, W.J., Rao, M.S., and Zeng, X. 2006. Assessing self-renewal and differentiation in human embryonic stem cell lines. Stem Cells 24:516-530. Carpenter, A.E., Jones, T.R., Lamprecht, M.R., Clarke, C., Kang, I.H., Friman, O., Guertin, D.A., Chang, J.H., Lindquist, R.A., Moffat, J., Golland, P., and Sabatini, D.M. 2006. CellProfiler: Image analysis software for identifying and quantifying cell phenotypes. Genome Biol. 7:R100. Chadwick, K., Wang, L., Li, L., Menendez, P., Murdoch, B., Rouleau, A., and Bhatia, M. 2003. Cytokines and BMP-4 promote hematopoietic differentiation of human embryonic stem cells. Blood 102:906-915. Chambers, I., Colby, D., Robertson, M., Nichols, J., Lee, S., Tweedie, S., and Smith, A. 2003. Functional expression Nanog c.o., a pluripotency sustaining factor in embryonic stem cells. Cell 113:643-655. Draper, J.S., Pigott, C., Thomson, J.A., and Andrews, P.W. 2002. Surface antigens of human embryonic stem cells: Changes upon differentiation in culture. J. Anat. 200:249-258. Enver, T., Soneji, S., Joshi, C., Brown, J., Iborra, F., Orntoft, T., Thykjaer, T., Maltby, E., Smith, K., Dawud, R.A., Jones, M., Matin, M., Gokhale, P., Draper, J., and Andrews, P.W. 2005. Cellular differentiation hierarchies in normal and culture-adapted human embryonic stem cells. Hum. Mol. Genet. 14:3129-3140. Forsyth, N.R., Musio, A., Vezzoni, P., Hamish, A., Simpson, R.W., Noble, B.S., and McWhir, J. 2006. Physiologic oxygen enhances human embryonic stem cell clonal recovery and reduces chromosomal abnormalities. Cloning Stem Cells 8:16-23. Gallagher, S.R. and Desjardins, P.R. 2006. Quantitation of DNA and RNA with absorption and fluorescence spectroscopy. Curr. Protoc. Mol. Biol. 76:A.3D.1-A.3D.21. He, J. and Landau, N.R. 1995. Use of a novel human immunodeficiency virus type I reporter virus expressing human placental alkaline phosphatase to detect an alternative viral receptor. J. Virol. 69:4587-4592.

Avilion, A.A., Nicolis, S.K., Pevny, L.H., Perez, L., Vivian, N., and Lovell-Badge, R. 2003. Multipotent cell lineages in early mouse development depend on SOX2 function. Genes Dev. 17:126140.

Henderson, J.K., Draper, J.S., Baillie, H.S., Fishel, S., Thomson, J.A., Moore, H., and Andrews, P.W. 2002. Preimplantation human embryos and embryonic stem cells show comparable expression of stage-specific embryonic antigens. Stem Cells 20:329-337.

Boiani, M. and Sch¨oler, H.R. 2005. Regulatory networks in embryo-derived pluripotent stem cells. Nat. Rev. Mol. Cell Biol. 6:872884.

Ji, L., Allen-Hoffmann, B.L., Juan Pablo, J.D., and Palecek, S.P. 2006. Generation and differentiation of human embryonic stem cell-derived keratinocyte precursors. Tissue Eng. 12:665-679.

Boyer, L.A., Lee, T.I., Cole, M.F., Johnstone, S.E., Levine, S.S., Zucker, J.P., Guenther, M.G., Kumar, R.M., Murray, H.L., Jenner, R.G., Gifford, D.K., Melton, D.A., Jaenisch, R., and Young, R.A. 2005. Core transcriptional regulatory circuitry in human embryonic stem cells. Cell 122:947-956.

Josephson, R., Ording, C.J., Liu, Y., Shin, S., Lakshmipathy, U., Toumadje, A., Love, B., Chesnut, J.D., Andrews, P.W., Rao, M.S., and Auerbach, J.M. 2007. Qualification of embryonal carcinoma 2102Ep as a reference for human embryonic stem cell research. Stem Cells 25:437-446.

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Lutfalla, G. and Uze, G. 2006. Performing quantitative reverse-transcribed polymerase chain reaction experiments. Methods Enzymol. 410:386400. Mitsui, K., Tokuzawa, Y., Itoh, H., Segawa, K., Murakami, M., Takahashi, K., Maruyama, M., Maeda, M., and Yamanaka, S. 2003. The homeoprotein Nanog is required for maintenance of pluripotency in mouse epiblast and ES cells. Cell 113:631-642. Niwa, H., Miyazaki, J., and Smith, A.G. 2000. Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat. Genet. 24:372-376. Oka, M., Tagoku, K., Russell, T.L., Nakano, Y., Hamazaki, T., Meyer, E.M., Yokota, T., and Terada, N. 2002. CD9 is associated with leukemia inhibitory factor-mediated maintenance of embryonic stem cells. Mol. Biol. Cell 13:1274-1281. Pera, M.F., Blasco-Lafita, M.J., Cooper, S., Mason, M., Mills, J., and Monaghan, P. 1988. Analysis of cell-differentiation lineage in human teratomas using new monoclonal antibodies to cytostructural antigens of embryonal carcinoma cells. Differentiation 39:139-149. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399404. Riedy, M.C., Muirhead, K.A., Jensen, C.P., and Stewart, C.C. 1991. Use of a photolabeling technique to identify nonviable cells in fixed homologous or heterologous cell populations. Cytometry 12:133-139. Rosner, M.H., Vigano, M.A., Ozato, K., Timmons, P.M., Poirier, F., Rigby, P.W., and Staudt, L.M. 1990. A POU-domain transcription factor in early stem cells and germ cells of the mammalian embryo. Nature 345:686-692.

Schatten, G., Smith, J., Navara, C., Park, J., and Pedersen, R. 2005. Culture of human embryonic stem cells. Nat. Methods 2:455-463. Schopperle, W.M. and Dewolf, W.C. 2006. The Tra1-60 and Tra-1-81 human pluripotent stem cell markers are expressed on podocaluxin in embryonal carcinoma. Stem Cells 25:723-730. Segev, H., Kenyagin-Karsenti, D., Fishman, B., Gerecht-Nir, S., Ziskind, A., Amit, M., Coleman, R., and Itskovitz-Eldor, J. 2005. Molecular analysis of cardiomyocytes derived from human embryonic stem cells. Dev. Growth Differ. 47:295-306. Stewart, M.H., Boss´e, M., Chadwick, K., Menendez, P., Bendall, S.C., and Bhatia, M. 2006. Clonal isolation of hESCs reveals heterogeneity within the pluripotent stem cell compartment. Nat. Methods 3:807-815. Takeda, J., Seino, S., and Bell, G.I. 1992. Human Oct3 gene family: cDNA sequences, alternative splicing, gene organization, chromosomal location, and expression at low levels in adult tissues. Nucl. Acids Res. 20:4613-4620. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Toh, W.S., Yang, Z., Liu, H., Heng, B.C., Lee, E.H., and Cao, T. 2007. Effects of culture conditions and BMP2 on extent of chondrogenesis from human embryonic stem cells. Stem Cells 25:950960. Trounson, A. 2006. The production and directed differentiation of human embryonic stem cells. Endocr. Rev. 27:208-219. Wong, M.L. and Medrano, J.F. 2005. Real-time PCR for mRNA quantitation. Biotechniques 39:75-85. Xu, X., Kahan, B., Forgianni, A., Jing, P., Jacobson, L., Browning, V., Treff, N., and Odorico, J. 2006. Endoderm and pancreatic islet lineage differentiation from human embryonic stem cells. Cloning Stem Cells 8:96-107.

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Isolation of Human Embryonic Stem Cell–Derived Teratomas for the Assessment of Pluripotency

UNIT 1B.4

Karin Gertow,1,7 Stefan Przyborski,2 Jeanne F. Loring,3 Jonathan M. Auerbach,4 Olga Epifano,4 Timo Otonkoski,5 Ivan Damjanov,6 and Lars 1,8 ¨ Ahrlund-Richter 1

Department of Laboratory Medicine, Clinical Research Center, Unit for Molecular Embryology, Karolinska Institute, Sweden 2 School of Biological and Biomedical Science, University of Durham, Durham, United Kingdom 3 Burnham Institute for Medical Research, LaJolla, California 4 GlobalStem Inc., Rockville, Maryland 5 Hospital for Children and Adolescents and the Biomedicum Stem Cell Center, University of Helsinki, Finland 6 Department of Pathology, The University of Kansas, School of Medicine, Kansas City 7 Monash Immunology and Stem Cell Laboratories, Monash University, Australia 8 Department of Woman and Child Health, Karolinska Institute, Stocholm, Sweden.

ABSTRACT This unit describes protocols on how to assess the developmental potency of human embryonic stem cells (hESCs) by performing xenografting into immunodeficient mice to induce teratoma formation. hESCs can be injected under the testis capsule, or alternatively into the kidney or subcutaneously. Teratomas that develop from grafted hESCs are surgically removed, fixed in formaldehyde, and paraffin embedded. The tissues in the teratoma are analyzed histologically to determine whether the hESCs are pluripotent and form tissues derived from of all three embryonic germ layers (ectoderm, mesoderm, and endoderm). Teratomas can also be fixed in Bouin’s or cryosectioned for analysis, and they can be analyzed by immunohistochemistry for tissue markers. Methods for these C procedures are included in this unit. Curr. Protoc. Stem Cell Biol. 3:1B.4.1-1B.4.29.
2007 by John Wiley & Sons, Inc. Keywords: human embryonic stem cells r pluripotency r teratoma r immunodeficient mice

INTRODUCTION Embryonic stem cells (ESCs) derived from the inner cell mass of the mammalian blastocyst have enormous potential for improvements in modern biology and medicine (Keller, 2005). The capacity of ESCs for multi-lineage differentiation can be demonstrated in culture; both mouse and human ESCs spontaneously produce a wide range of cell types derived from all three embryonic germ layers: ectoderm, mesoderm, and endoderm. For mouse ESCs, definitive proof of their developmental pluripotency is routinely obtained by demonstrating their ability to develop into all cell types, including germ cells, in chimeras produced by mingling ESCs with mouse blastomeres or blastocysts (Evans and Kaufman, 1981; Martin, 1981). Since human ESCs (hESCs) cannot be subjected to the same definitive test used to prove developmental pluripotency of mouse ESCs, the most rigorous method available for testing hESC is to inject them into immunodeficient mice and analyze histologically the composition of teratomas formed from the cells (Thomson et al., 1998). Teratomas Current Protocols in Stem Cell Biology 1B.4.1-1B.4.29 Published online October 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b04s3 C 2007 John Wiley & Sons, Inc. Copyright


Embryonic and Extraembryonic Stem Cells

1B.4.1 Supplement 3

Table 1B.4.1 Summary of Protocols

Protocol function

Advantages

Disadvantages

Surgery is straightforward and testis easily accessible

Teratomas occasionally form more cystic type structures— reason unknown

Teratoma growth is easily detected by palpation

Large tumor growth can interfere with bladder function and blockage

Transplantation site Testis (Basic Protocol 1)

Teratomas most often confined within the — capsule and contain well differentiated tissues Kidney (Alternate Protocol 1)

Well-vascularized location

Surgery is more complicated

Tumors most often confined within the Difficult to follow tumor growth by palpation capsule and contain well-differentiated tissues Subcutaneous (Alternate Protocol 2)

—

Measurement by calipers is not possible

Straightforward method of implanting cells that does not require surgery or anesthesia

Can be a poorly vascularized location

Tumor growth is easily recognizable and can be monitored by caliper measurements

Large tumors become pronounced and skin covering may become eroded due to surface abrasion

Least invasive site and less likely to compromise the welfare of the host

—

Multiple graft sites can be used

—

Avoids enzyme treatment and washing

Time consuming

Allow for selection of colonies with desired morphology

Difficult to quantitate number of cells accurately

Aggregates of cells are more likely to be retained at the transplant site

Large numbers of cells/aggregates may be difficult to transplant

Preparation of hESC Mechanical (Support Protocol 1)

Enzymatic (Support Protocol 1)

Allows harvest of a precise or larger numbers Cells spread more easily at the transplantation site of cells (>105 )

Tissue preservation Paraformaldehyde (Basic Protocol 2)

Preserves tissue morphology well

Good for immunohistochemistry

Bouin’s fixative (Alternate Protocol 3)

Preserves tissue morphology well and gives excellent preservation of nuclei and chromosomes

Suboptimal for immunohistochemistry

Cryopreservation (Alternate Protocol 4)

Can be used for DNA or RNA studies

Suboptimal for tissue morphology

Isolation of hESC-Derived Teratomas

derived from hESCs contain demarcated areas containing differentiated cells and tissues representing all three embryonic germ layers. These tissues are in many ways reminiscent of structures found during early organogenesis (Reubinoff et al., 2000; Richards et al., 2002; Gertow et al., 2004). The ability to give rise to well-differentiated teratomas is a defining feature of pluripotent ESCs and is a key measure of the stem cells’ abilities to develop into various types of tissues. In spite of the wealth of information provided by this test, it has surprisingly not been rigorously applied as a standard assay for characterizing hESC lines.

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Teratomas are most often produced in immunodeficient mice by injecting hESCs beneath the testicular or kidney capsule (Thomson et al., 1998; Reubinoff et al., 2000; Lanzendorf et al., 2001; Richards et al., 2002; Hovatta et al., 2003; Mitalipova et al., 2003; Park et al., 2003; Heins et al., 2004; Pickering et al., 2005). Cells can also be injected intramuscularly (Tzukerman et al., 2003; Li et al., 2004), subcutaneously (Levenberg et al., 2002), or in the liver (Cooke et al., 2006). In general, injection sites at which the teratoma growth can be monitored by external examination, without interference with general well-being seem to be preferred to approaches that demand surgical exploration. Except for the liver, where grafted cells produce large tumors made of immature cells, all other sites seem to encourage development of differentiated tissue, often identifiable as derivatives from all three germ layers (Cooke et al., 2006). This unit provides protocols that can be used to demonstrate pluripotency of hESC preparations and to assess their developmental potential by inducing teratomas in immunodeficient mice (summarized in Table 1B.4.1).

STRATEGIC PLANNING Choice of Injection Site Depending on the type of question to be addressed, different sites of injection may be appropriate. Here the authors describe protocols for hESC grafting under the testis (Basic Protocol 1) or kidney capsule (Alternate Protocol 1) or for subcutaneous injection (Alternate Protocol 2). Type of Analysis Depending on the type of information desired, different types of analysis may be applied. Here the authors describe protocols for an analysis of tissue morphology and marker analysis using immunohistochemistry. In addition they have included a protocol for an RNA expression analysis performed on hESC teratoma tissue samples. The choice of markers will influence the protocol necessary for the preparation of the tissue. Choice of Recipient Mouse Host Studies have been reported using severe combined immunodeficient (SCID) mice (Reubinoff et al., 2000; Stojkovic et al., 2004a), or nude (nu) mice (Mikkola et al., 2006; Yao et al., 2006). Alternatively, combinations of these mutations with either beige (bg; Thomson et al., 1998; Gertow et al., 2004) or nonobese diabetic (NOD) have been used (Ito et al., 2002). General Advice Immunodeficient mice must be housed using barrier rearing facilities (e.g., isolator or separately ventilated cages). When handling immunodeficient mice one must keep in mind that these animals have an increased susceptibility to infections. Mice must be fully healthy and not stressed. Avoid surgery on mice the same day as the cages have been cleaned, as this causes stress and makes the mice less responsive to anesthesia. Use a minimum of five mice for testing each cell type injected.

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NOTE: All protocols using hESCs and irradiation of feeder cells must be reviewed under existing mandates and by following the rules and regulations set by local committees with jurisdiction over research on human tissues, biosafety, and radiation. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations for the care and use of laboratory animals. NOTE: Each animal will receive only one injection of cells in a particular site. BASIC PROTOCOL 1

INJECTION OF hESC UNDER THE TESTIS CAPSULE IN IMMUNODEFICIENT MICE This protocol describes the surgical procedure for injecting hESCs into an immunodeficient mouse model, for the purpose of demonstrating a pluripotent capacity of the grafted cells. The testis is not a vital organ, it is relatively easy to access, and the teratoma growth can, at least partially, be monitored by external examination.

Materials hESCs (see Support Protocol 1) Mice (immunodeficient; either from immunosuppressive treatment, or genetic mutation; see Strategic Planning) 70% ethanol 0.015 mg/ml Temgesic (Buprenorphinum) Stereomicroscope for harvesting hESCs in the animal surgery room (optional depending on procedure used for the harvest of hESCs; see Support Protocol 1) Sterile paper tissue Electric clippers Sterile drapes 2 curved forceps Small surgical scissors Dissecting microscope 1-ml syringe (e.g., U-100 Micro-Fine 12.7-mm; Becton Dickinson), or a Hamilton syringe Needle holder Culture dish Resorbable sutures (e.g., Ethicon, Vicryl V422 4-0) 9-mm stainless steel wound clips (autoclips from MikRon Precision) Clip applier Additional reagents and equipment for anesthetizing the recipient mouse (Support Protocol 2) NOTE: Protocols for surgical opening of the abdomen require the use of sterile instruments, surgical gloves, and aseptic procedures to minimize the risk of post-surgical infection.

Prepare cells and mice 1. Prepare the hESCs (Support Protocol 1) and anesthetize the recipient mouse (Support Protocol 2). Open the abdomen 2. Clean the anterior wall of the abdomen with paper tissue soaked in 70% ethanol. Isolation of hESC-Derived Teratomas

Do not wet the animal directly with excess amounts of ethanol, since this may cool the animal.

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Figure 1B.4.1 Instrument set up and schematic procedure for the intra testicular implantation. (A) The anesthesia unit comprised of the (1) Univentor × 2, (2) induction chamber, and (3) maintenance mask. (B) Position of the mouse in the maintenance mask and the incision along linea alba (LA). (C,D) hESC injection under the testis capsule. T = testis; F = fat tissue; E = epididymis (E) Effect from injecting too large a volume under the testis capsule.

3. Using scissors and/or electric clippers, shave an area on the lower part of the abdomen and rinse with 70% ethanol. Drape the area with sterile drapes. This helps prevent hair from entering the surgical field and provides a clean area on which to lay the testes in steps 7 to 11 below.

4. With forceps gently pull up the skin on the lower part of the abdomen and with scissors make a 1to 2-mm cut vertically along the midline (linea alba; see Fig. 1B.4.1B). Using the linea alba incision reduces the risk of bleeding from inadvertently cut vessels.

5. Next, cut 5 to 10 mm further from the first cut along the linea alba towards the head. The size of the incision should allow for the testis and attached fatty tissue to pass through easily but without being unnecessarily large.

6. With the forceps grip the edge of the wound on one side and carefully separate the outer skin from the peritoneum by placing closed scissors under the skin and gently opening the scissors. Repeat on the other side. This will force the skin to detach from the peritoneum and makes it easier to stitch the peritoneal sutures while avoiding the skin.

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7. Change the grip of the forceps, carefully pull the peritoneum and cut the peritoneum along the linea alba. At this stage it is vital to avoid cutting any internal organs. The peritoneal incision should be shorter/smaller than that of the skin; otherwise it may be difficult to close the peritoneum (step 14 below).

8. Grip the edge of the peritoneum with one pair of forceps and with a second pair of forceps reach in down towards the right hip and very gently pull up the fatty tissue along with the attached testis. Position the testis outside of the abdomen on the sterile drape (Fig. 1B.4.1C). This step requires special attention when gripping the internal fat tissue in order to avoid damage! It is easy to unintentionally grip intestines instead of the fatty tissue. Note the fatty tissue is white while intestines are beige in color. Although this protocol recommends using the right testis, either side can be used, but it is important to routinely implant on a particular side. It is possible to use the same surgical instruments on more than one animal. If care is used to maintain asepsis of surgical instruments, they may be used for a maximum of five animals.

Inject hESC 9. Set up a dissecting microscope in the operating area. Perform initial cell injections using low magnification with a dissecting stereomicroscope. Although the injections can also be performed without a microscope, it is advised to use a microscope until sufficient surgical skill is attained.

10. Puncture the testicular capsule, close to where the testis is attached at the proximal base, with the needle of the injection syringe. Position the needle so the cells can be injected into the center of the testis and inject the hESCs (20 to 30 µl; see Support Protocol 1, steps 3 to 4; Fig. 1B.4.1C to 1B.4.1E). When puncturing the testicular capsule, care must be taken to avoid puncturing vessels since this can cause bleeding and unnecessary discomfort. The volume injected under the testes capsule should not exceed 30 µl, or it may cause rupture of the capsule (Fig. 1B.4.1E).

11. Withdraw the syringe slowly in order to avoid capsule rupture or reflux of the cells from the injected organ. 12. Aspirate the syringe up and down several times in a culture dish containing culture medium. This will verify that the cells were injected and not stuck inside the syringe. If the injection failed you may have a chance to perform a second injection at the same site, depending on the status of the animal.

13. Next, place the testis and fatty tissue back in the correct position in the abdominal cavity by gripping the fatty tissue using one pair of forceps and with a second pair of forceps holding and separating the peritoneal wall in order to push the testis and fat pad into position. Take care not to apply any pressure on the testis, since this may cause the cells to flow out or the capsule to rupture.

Close the incision 14. Close the peritoneum using resorbable sutures, commonly 2 to 3 stitches in total. Isolation of hESC-Derived Teratomas

It is critical to avoid placing the sutures through the intestines and the underlying fatty tissue.

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15. Close the skin incision by holding the two sides of the cut together with a pair of forceps and staple with wound clips using a clip applier to seal the skin. 16. To comply with policies on minimizing pain and distress in laboratory animals, use analgesics in the post-surgical period. For example, inject 0.05 to 0.1 mg/kg Temgesic (Buprenorphinum, 0.015 mg/ml) per mouse intraperitoneally.

Inspect post-surgically 17. While the mouse is still anesthetized, label the mouse (e.g., ear marking). 18. Move the animal to a warm, dry area and monitor its recovery before returning it to the original cage. Avoid placing mice from different cages together. They will fight as soon as they recover from anesthesia.

19. Mark cages with a label containing the details of the procedure.

Provide follow-up care 20. Carefully follow the health status of the mice daily the first week after surgery. 21. Remove wound clips when the wound is fully healed (typically within 2 weeks). 22. When an hESC teratoma is palpable, check the animals every second to third day until the end of the experiment. Normally the animals show no signs of distress from hESC teratoma growth unless the growth is allowed to progress beyond the size recommended for this protocol. Always sacrifice mice that are not fully healthy. Occasionally, the testis is not re-localized to the scrotum and development of hESC teratoma in the abdomen can be difficult to detect by palpitation. Besides the health status of the animal, the end point is dependent on the experimental question and the hESC dose injected. 104 to 105 cells injected generally generate a teratoma within 6 to 10 weeks, and may be grown further up to 12 weeks. Higher doses, >106 cells, can generally be harvested within 6 weeks. The end point for the hESC teratoma growth should be determined by: A predefined time after implantation Palpable growth beyond a certain size Health status of the animal. The animal’s health status must at all times be the first priority! Monitoring the animal’s weight is a useful indicator of tumor progression, especially if a growing xenograft is not palpable.

INJECTION OF hESC UNDER THE KIDNEY CAPSULE This protocol describes an alternative injection under the kidney capsule, an approach commonly used by many research groups and proven very effective in demonstrating the pluripotent capacity of the injected cells.

ALTERNATE PROTOCOL 1

For materials see Basic Protocol 1.

Open the abdomen 1. Perform steps 1 to 7 of Basic Protocol 1. See general advice given in Basic Protocol 1, steps 1 to 7.

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2. Instead of reaching for the testes fat tissue, localize the kidney and without repositioning the organ proceed as described below. Alternatively, place the mouse on its right side, and place a microcentrifuge tube underneath, so that the left kidney site is protruding. Make the initial incision just above the kidney, beneath the spleen. Once the kidney is exposed, carefully avoid touching it, by applying some pressure and pulling from the connective tissue surrounding the kidney.

Inject hESC 3. Inject the cells under the kidney capsule (Basic Protocol 1, steps 9 to 11). Preferably, inject cells on the lateral side of the kidney to avoid damage to any major blood vessels. Insert the needle at an acute angle to enable the release of cells into the sub-capsular region of the organ a few millimeters lateral to the needle entry point. Subsequent to injection of the cell suspension, slowly remove the needle to minimize cell loss and tissue damage. It is essential that the volume of the cell suspension is kept to a minimum and no air is introduced during the injection procedure to avoid disruption to the internal structure of the kidney. Alternatively, hESCs can be delivered via a capillary attached to a mouth pipet with a filter. A small incision is made with micro-scissors into the kidney capsule. This has to be done very superficially, with no bleeding. A sterile glass rod of the same thickness as the capillary is then used to make a small “pocket” for the cells immediately underneath the capsule. The rod is removed and immediately replaced by the capillary of the mouth pipet, through which the cells are gently blown into the pocket.

4. Proceed with Basic Protocol 1, steps 12 to 22, to close the incision, inspect the animals post-surgically, and provide follow-up care. ALTERNATE PROTOCOL 2

SUBCUTANEOUS INJECTION OF hESC Subcutaneous injection of hESCs is the least invasive injection route for demonstrating a pluripotent capacity of the injected cells. However, the cells are not retained at the injection site as well as when injected into an encapsulated organ. Thus, the efficiency of teratoma generation is lower. The benefits of subcutaneous injection are that the growth can be easily followed by external palpation and if a scaffold is to be used, this is possibly the site of choice.

Additional Materials (see Basic Protocol 1) Phosphate buffered saline, calcium- and magnesium-free (CMF-PBS) 21-G needle NOTE: Subcutaneous injection of cells does not require anesthesia. 1. Inject ∼5 × 105 cells in 50 µl CMF-PBS using a 21-G needle subcutaneously into the flank. IMPORTANT NOTE: It is important to keep the injection volume low (<100 µl). Following injection of the hESCs, it is advisable to aspirate the syringe up and down a couple of times in a culture dish containing medium to verify that the cells are injected and not stuck inside the syringe.

Isolation of hESC-Derived Teratomas

2. For the follow-up care of the hESC-injected mice, follow steps described in Basic Protocol 1. Maintain mice for up to 12 weeks, and monitor animal welfare and tumor progression regularly. When a teratoma is identified, record location and size of the tumor (using measuring calipers) and monitor the weight of the animal. A teratoma is usually first identified as a small palpable mass beneath the skin near to the transplantation site.

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PREPARATION OF hESC FOR INJECTION This protocol describes the major steps to be considered when preparing hESCs for injection. Standardized protocols for the preparation of hESCs will help to optimize the teratoma formation and to make comparisons between experiments possible.

SUPPORT PROTOCOL 1

Materials Cultures of hESC Enzyme for harvesting Dissecting microscope or stereomicroscope 1-ml insulin syringe Mouth pipet Thick needle Equipment for enzymatic splitting Stereomicroscope Centrifuge Select cells 1. Passage the cells, using a standard culture protocol, 2 to 3 days before injecting them into mice. The cells should be in logarithmic growth phase and not confluent when transplanted in vivo. It is recommended to use cells cultured under standard conditions and at a passage number equivalent to that used for other experiments. Ideally, use single-well culture dishes/plates in order to avoid unnecessary time spent outside the incubator during harvest and injection of the cells in adjacent wells. For instance, use center-well organ culture dishes (60 × 15–mm) or cell culture dishes (60 × 15–mm). Avoid transporting the cells, unless good culture conditions can be maintained during the transport. It is the authors’ experience that hESCs maintained for transport for >1 hr at room temperature do not engraft as readily. When comparing different hESC lines, take care to grow them under the same culture conditions and transplant them at a similar passage number. Small variations in culture medium, especially in serum or serum-free supplement, can dramatically alter cell differentiation and lead to differences in teratoma development. This makes characterization and comparison of different cell lines particularly problematic.

2. On the day of injection, thoroughly evaluate the morphological quality of the cells under a microscope. Transport the culture dish with the cells to the operation room, taking care to avoid metabolic stress to the cells and to maintain sterility. It is highly recommended to keep the cells in a cell culture incubator in the operation room while working surgically with the mice. It is important to handle the cells as little as possible following harvest from the culture dish and also to keep the time prior to injection as short as possible. To avoid the need for enzyme treatment and washing procedures including centrifugation, the authors have found it useful to harvest the cells in culture medium mechanically immediately prior to injection. There is no need to remove mitotically inactivated feeder cells from the hESC preparation.

Harvest hESCs Mechanical harvesting of hESC colonies 3a. Under a dissecting microscope, select undifferentiated colonies of good morphology. Using a 1-ml insulin syringe, cut around the colony (harvest them one by one), gently free the cells from the dish by scraping using the syringe. With the same syringe, aspirate the colony in as small a volume as possible (this takes some training). Depending on the size of the colonies, 1 to 3 colonies will be equivalent to 104 cells. Larger colonies should be broken up into smaller pieces in order not to clog the needle. Make sure not to aspirate >30 µl (0.03 ml) total volume.

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Figure 1B.4.2

Undifferentiated hESC colonies, each containing ∼3000 cells.

The volume injected under the testis capsule should not exceed this volume, or it may cause rupture of the testicular capsule. It may be helpful to transfer the colonies to a small drop and collect cells/colony fragments from the drop to minimize the volume of fluid to be injected. With some experience in the culturing of the individual hESC line it is possible to predict fairly well the cell numbers in individual colonies (Fig. 1B.4.2).

3b. Alternatively, collect the cells (colonies) under a stereomicroscope using a mouth pipet and a thick needle, so that the excess fluid is blown out but the cell clumps remain in the capillary, held by the needle. This only works for clumps of cells, not for a single-cell suspension.

Enzymatic harvesting of HESC 3c. When injection of precise or larger numbers of cells (>105 ) is essential, harvest using enzyme and disperse into a single-cell suspension using your standard protocol for enzymatic splitting of hESCs and count the cells. We recommend using an enzyme not requiring inactivation with an inhibitor, such as TrypLE or dispase. However, it is more important not to alter your standard passaging protocol.

4c. It is important to remove or inactivate residual enzyme. Concentrate the cells by centrifuging 3 min at 450 × g, 4◦ C, immediately before injection and add minimal injection volume, in order to maintain smallest possible injection volume. Generally, the use of the lowest dose for engraftment (in the authors’ experience 103 to 105 per inoculum) is recommended, although the actual number of cells needed for transplantation must be decided by the experimental situation. Isolation of hESC-Derived Teratomas

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ANESTHESIA FOR THE MOUSE Protocols that include opening of the abdomen require anesthesia. Mice lose body temperature and fluid fast, especially when they are opened during surgery. Therefore, the shorter time the animal needs to be anesthetized the better. To prevent cooling a heating blanket or lamp may be used. It may also be necessary to replace fluid with 10 to 15 ml/kg warmed 0.9% saline administered subcutaneously. Generally, if the entire procedure from anesthesia to removal of the anesthetics takes 10 min or less, it is enough to post-surgically move the animal to a warm, dry area and monitor it during recovery. If the entire procedure takes longer (than 10 min), the above actions should be taken. For further details on standard surgical procedures, see Waynforth and Flecknell (1999).

SUPPORT PROTOCOL 2

Materials Recipient animals Anesthetics (in compliance with local guidelines for major surgery): e.g., isoflurane (1-chloro-2.2.2-trifluoroethyl difluoromethyl ether) Ophthalmic ointment or artificial tears Hypnorm (fentanyl/fluanisone) Anesthesia unit: induction chamber (0.8-liter) and maintenance mask (e.g., Univentor 400; http://www.univentor.com) Surgical tape Anesthetize with isoflurane inhalation 1a. Before starting, make a presurgical evaluation of the recipient animals. Select only healthy animals. Also, make sure all equipment is tested and functional. 2a. Set the induction chamber at 4% isoflurane and a flow of air at 400 ml/min. IMPORTANT NOTE: Since in these protocols an inhaled anesthetic is used, the surgery must be performed in a well-ventilated area. Properties of isoflurane: (1) Inhalation agent that produces a rapid induction and rapid recovery. (2) Nonflammable and nonexplosive. (3) Irritant to airways. (4) Low biotransformation/does not induce liver enzymes/removed by exhalation. (5) Can cause vasodilation resulting in hypotension and tachycardia.

3a. Set the maintenance mask at 1.7% to 3% isoflurane and a flow of air at 200 to 250 ml/min. These concentrations are only recommendations and must be decided by the experimenter. The fresh gas flow rates should exceed three times the animals’ minute volume. This protocol uses two separate anesthesia units: one for the induction chamber and one for the maintenance gas mask (Fig 1B.4.1A). It is also possible to use only one anesthesia unit together with a gas routing switch (cat. no. 8433005, Univentor). Always ensure that the system is loaded with sufficient volumes of isoflurane before initiating anesthesia on each animal. The volume needed depends on concentrations/flow and time used for the surgery. Estimate 10 to 15 min per animal for a skilled surgeon.

4a. Place the mouse in the induction chamber (Fig. 1B.4.1A). Avoid causing extra stress to the animal. A stressed animal will be difficult to anesthetize. Lift the mouse by holding it firmly at the base of the tail and allow the mouse to grip a surface (such as your hand) still holding a firm grip of the tail. Only open the chamber immediately before the animal is moved into it; this avoids allowing the gas to escape, lowering the concentration and causing unnecessary stress to the animal.

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5a. When the mouse is still and breathing calmly, quickly move the animal to the maintenance mask and position it for surgery; i.e., with the abdomen upwards and the head pointing away from you (Fig. 1B.4.1B). Use surgical tape to gently secure the position of the animal. The animals awake quickly after removing the anesthesia so this step must be swift. Make sure the animal’s head is properly placed in the maintenance mask. Taping is done to avoid repositioning the animal away from the maintenance mask, which risks the animal awakening. Note that the tape should not be firmly stretched.

6a. Assess the depth of anesthesia: pinch a digit or interdigital skin with a forceps after 10 to 20 sec in the maintenance mask to test the pedal reflex. The mouse is fully anesthetized when this reflex disappears. If the animal reacts in any way, control the position in the maintenance mask and wait another 10 to 20 sec. Repeat the procedure. If there still is a reaction from pinching, most likely the gas concentrations need to be regulated.

7a. Place a drop of protective ophthalmic ointment or artificial tears on each eye to protect the cornea from drying. 8a. Monitor the color of the mucous membranes, the respiration, and the heart rate during the entire anesthesia and surgery procedures. If the animal awakens or shows any signs of reaction at any time during the surgery, it must immediately be euthanized.

Anesthetize using fentanyl/fluanisone (Hypnorm) 1b. Before starting, make a presurgical evaluation of the recipient animals. Use only healthy animals. Also, make sure all equipment is tested and functional. Avoid causing extra stress to the animal. A stressed animal will be difficult to anesthetize.

2b. Inject 0.65 ml/kg hypnorm, i.p. or 0.4 ml/kg i.m. to induce surgical anesthesia For properties of Hypnorm, see http://www.vetapharma.co.uk/.

3b. Follow steps 6a to 8a above. BASIC PROTOCOL 2

EXCISION AND FIXATION FOR PARAFFIN EMBEDDING OF THE hESC TERATOMA This protocol describes how to excise the teratoma tissue and how to process it by fixation. The fixative used in this protocol cross-links proteins, which helps to preserve the tissue morphology, a prerequisite for histological analysis. NOTE: Euthanizing an animal must be painless and death should be induced rapidly while the animal is unconscious. Training of personnel and veterinarian supervision of the first procedures may be required. NOTE: Depending on how tissues are to be analyzed, the way they are processed differs. Alternate Protocol 3 describes fixation using immersion in Bouin’s solution and Alternate Protocol 4 describes preparation for cryosectioning.

Materials

Isolation of hESC-Derived Teratomas

Mouse with teratoma 70% ethanol Sodium pentobarbitone (for perfusion fixation experiments) 4% (w/v) buffered formaldehyde or paraformaldehyde (PFA) in saline Paraffin wax

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Sterile paper tissue Scissors Forceps curved and straight 10-cm dish Razor blade 50-ml tubes Additional reagents and equipment for euthanizing the mouse (Donovan and Brown, 2006) and fixation, embedding, and sectioning tissues (http://home.primus.com.au/royellis/histo.html or Hofman, 2002) CAUTION: Formaldehyde is an irritant; avoid skin contact and inhalation of its vapors.

For regular fixation 1a. Euthanize the mouse (Donovan and Brown, 2006) in compliance with local regulations. This may be accomplished by dislocating the cervical vertebrae by stretching the animal and rotating the neck. This way the spinal cord is disrupted and nerve impulses to the vital organs are no longer transmitted.

2a. Swab the animal with paper tissues soaked in 70% ethanol to clean the abdominal surface.

For perfusion fixation 1b. Administer a lethal dose of anesthetic (e.g., 100 mg/kg sodium pentobarbitone, i.p.). 2b. Using scissors, open the thoracic cavity to expose the heart. 3b. Introduce saline and fixative solutions into the left ventricle, allow to circulate and exit via the excised right atrium. See http://www.chemicon.com/techsupp/Protocol/perfusion.asp for details.

4. Open the abdomen using scissors and forceps. 5. Locate and cut the testis/teratoma loose from fatty tissue and seminiferous tubules. The testis/teratoma is commonly located in the lower abdominal part where the untreated testis is also normally located. Occasionally, the teratoma can be found located higher up in the abdominal cavity. The testis/teratoma should appear as a free and well-defined encapsulated structure. If alternative sites of injection are used, excise the teratoma from the appropriate area.

6. Place the tissue in a 10-cm dish and cut it into halves (or sections of no more than 6 to 8 mm in thickness) using a razor blade. Do not use a scalpel since this tends to cause more damage to the tissue.

7. Preserve surgically excised tumor tissues by submersion in 4% (w/v) formaldehyde in a large enough container, such as a 50-ml tube, to ensure sufficient fixation. Incubate at 4◦ C for 12 to 24 hr, with agitation to allow the fixative to reach all tissue surfaces. Fixation of thicker tissues may be sub-optimal and affect the quality of tissue histology since formaldehyde penetrates only slowly into the tissues. Avoid fixing the intact teratoma since the fixative solution penetrates the capsule ineffectively. Moreover, histological sections from the central parts of the hESC teratomas generally provide a better overview than tissue in the outer rim of the capsule.

8. Perform a necropsy of the animal, looking for detectable changes to other tissues and/or organs.

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9. After fixation, discard the fixative (in compliance with local regulations) and replace with 70% ethanol. Keep tubes at 4◦ C until paraffin embedded according to standard procedures (not more than 4 weeks). See http://home.primus.com.au/ royellis/histo.html or Hofman, 2002 for detailed protocols on fixation, embedding, and sectioning. 10. Process the tissues by dehydration through serial ethanol solutions and clearing followed by embedding in paraffin wax. Dehydration is usually initiated in 60% to 70% ethanol, progressing through 90% to 95% ethanol, then two or three changes of absolute ethanol before proceeding to the clearing stage. Clearing is the transition step between dehydration and infiltration with the embedding medium, often performed in Histoclear or xylene. Paraffin-embedded specimens can be serially sectioned (5- to 8-µm) using a standard rotary microtome, mounted on microscope slides and counterstained as appropriate. ALTERNATE PROTOCOL 3

BOUIN’S FIXATION OF TERATOMAS Bouin’s fixation offers good penetration and morphological/structural preservation of tissues. It is also a fixative compatible with a broad range of histological stains, although it is suboptimal for subsequent immunohistochemistry.

Additional Materials (also see Basic Protocol 2) Bouin’s fixative: 70% (v/v) saturated picric acid (Sigma); 25% (v/v) of 37% to 40% formaldehyde; 5% (v/v) glacial acetic acid (Sigma) Additional reagents and equipment for fixation, embedding, and sectioning tissues (http://home.primus.com.au/royellis/histo.html) and cryosectioning (Hofman, 2002) 1. Preserve surgically excised tumor tissues by submersion in Bouin’s fixative at room temperature for 12 to 24 hr. Tissues can subsequently be stored in 70% ethanol for a number of weeks prior to tissue processing and paraffin embedding.

2. Process tissues and embed in paraffin according to standard procedures. ALTERNATE PROTOCOL 4

TISSUE CRYOPRESERVATION AND PREPARATION FOR CRYO–MICROTOME SECTIONING For some enzyme and immunohistochemical analysis, fixation and paraffin embedding are not optimal and tissues may instead need to be frozen in liquid nitrogen using cryopreservative procedure. The following protocol is an alternative to Basic Protocol 2. Snap-frozen tissue can also be used for DNA or RNA studies.

Materials Freshly excised teratoma tissue Cryomount ( e.g., TissueTek OCT; Sakura) Liquid nitrogen

Isolation of hESC-Derived Teratomas

Razor blade Specimen holder/Cryomould Long forceps Cryotube Cryomicrotome SuperFrost+ slides

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CAUTION: Use protective gloves and glasses when handling liquid nitrogen. Use long forceps to place holders/cryotubes in liquid nitrogen. 1. Using a razor blade, section the freshly excised teratoma tissue into pieces no more than 6-mm thick. Thicker sections may freeze too slowly, causing ice crystallization. Therefore thinner sections are recommended.

2. Treat the tissue sections using one of the following alternatives: a. Mount each section in OCT TissueTek on a specimen holder (following the supplier’s instructions). Snap freeze (with tissue facing upwards) in liquid nitrogen. b. Place each section in a cryotube and snap-freeze in liquid nitrogen. Store at −70◦ C until mounted in OCT TissueTek. 3. Store at −70◦ C until sectioned on a cryomicrotome and collected on SuperFrost+ slides.

EVALUATION OF TISSUE FORMATION AND DEMONSTRATION OF THE PRESENCE OF EMBRYONIC GERM LAYERS IN THE hESC TERATOMA

BASIC PROTOCOL 3

Pluripotency of embryonic stem cells has been defined as the potential to differentiate into all three germ layers. Impaired development in a hESC teratoma from a given hESC line can be detected by the lack of all or some of the tissues representing a particular germ layer. Histological evaluation requires training and thus the authors strongly advise consulting a specialist. The protocol described here is intended to generate material useful for adequate analysis, but cannot replace the need for consulting pathology expertise. In this section some of the most readily identifiable tissues are illustrated. Immunohistochemistry may be required in some cases for a more precise identification and evaluation of tissues.

Teratoma Evaluation Helpful observations When excising the hESC teratoma the following observations are helpful for the evaluation of gross morphology: 1. Is the teratoma encapsulated or does it invade the surrounding tissues? Invasive growth is a feature of malignant tumors, whereas benign teratomas appear lobular and well circumscribed.

2. Assess color and consistency of the tumor on cross-section. Record whether the teratoma is predominantly solid or cystic. Dark regions commonly reflect hemorrhages or necrotic tissue (earlier harvest could prevent this).

3. Does the teratoma appear to contain fluid-filled (cystic) compartments? Histological evaluation is most often performed on paraffin-embedded sections of 5 to 7 µm, stained with hematoxylin and eosin (HE; Figs. 1B.4.3, 1B.4.4C,D and 1B.4.5A,C). Weigert’s stain can also be used (Fig. 1B.4.4A,B). This stain is traditionally used to stain for elastic fibers and myelin. It is not necessary to use Weigert’s stain in particular. Any general histological stain is acceptable, providing it enables the user to see the histological structure of the teratoma clearly. It is advisable to examine a minimum of 2 to 3 sections. Make sure sections are included from each embedded part of the teratoma. Fewer sections will not give a proper evaluation of the three-dimensional tissue.

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Figure 1B.4.3 Hematoxylin and eosin (HE) histology from formaldehyde-fixed teratoma tissues derived from human ES cells transplanted into the testis of SCID-beige mice and grown for 8 weeks. In (A) two different ectodermal epithelia are shown, (ep) of peridermal character and (p*) a pigmented epithelium possibly of retinal origin; (bn* and bn) intramembraneous bone formation and, (ca) cartilage are also seen. The structures (p*) and (bn*) from (A) are shown in higher magnification in (B). (C) Endodermal epithelia (en-ep) of intestinal character next to a fluid filled cystic (cy) region. (D) Striated muscle (mu) of mesodermal origin and a vessel (ve). Scale bars: (A) 150 µm; (B, C, D) 100 µm.

Questions to address Typical questions to address are as follows: 1. Does the section contain a mixture of different identifiable tissues? 2. Which tissues predominate and which are not present? 3. Are tissues derived from all three germ layers present?

hESC-derived teratoma tissues Examples of tissues commonly found in hESC-derived teratomas are shown in Figures 1B.4.3 through 1B.4.5: Mesoderm-derived tissues—bone and cartilage, striated and smooth muscle (Fig. 1B.4.3A,B,D; Fig. 1B.4.4A,B). Isolation of hESC-Derived Teratomas

Ectoderm-derived tissues—neuroepithelium/neuroectoderm, retinal-like pigmented epithelium, ganglia (Fig. 1B.4.3A,B; 1B.4.4B,C; 1B.4.5A,B).

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Figure 1B.4.4 Histological analysis of Bouin’s fixed teratoma tissues derived from human ES cells transplanted into the testis of severe combined immunodeficient mice and grown for 6 to 8 weeks. The images show an example of tissues representative of each germ layer. (A) Early stage of endochondral ossification, where cartilage (ca) transforms into bone. (B) Structure resembling wall of intestinal tract including surface epithelium (ep) lining crypt-like structures, underlying submucosa (mu), smooth muscle layer (sm), and neural ganglia (ng). (C) Neural ganglion (ng) with associated nerve fiber (nf). (D) Intestinal epithelium viewed under oil immersion using the ×100 objective lens showing high level of cellular detail, including a goblet cell (gc). Histological staining: Weigert’s (A,B) and hematoxylin and eosin (C,D). Scale bars: (A,C) 100 µm; (B) 150 µm; (D) 10 µm.

Endoderm-derived tissues—intestinal or bronchial epithelium (Fig. 1B.4.3C; 1B.4.4C,D) with mucin-producing goblet cells (Fig. 1B.4.4B,D), surrounded by layers of smooth muscle. Histological staining is useful for morphological visualization of tissues but is limited and is not specific to be informative about the true identity of a tissue or cell type, especially when dealing with tissues that are morphologically less distinct, such as pancreas and liver. As a general rule, most tissues may be identified histologically in HE-stained slides, but in some instances, additional immunostaining with antibodies to tissue-specific markers is needed for the positive identification of tissues. Some tissues are more commonly found (Fig. 1B.4.5); Table 1B.4.2 lists commonly observed tissues in hESC teratomas and primary antibodies used to detect them. It is important to keep in mind that few biomarkers show specific expression in only a single cell type; many molecules share expression patterns in different cells. Moreover, the expression of certain markers depends on the degree of tissue maturity and differentiation in the teratoma, stressing again the importance of expert pathohistological evaluation. If dealing with a teratoma that has not completely differentiated one could perform immunohistochemistry (see Basic Protocol 4; UNIT 1B.3) with biomarkers indicative of specific early germ layer differentiation, such as the nucleus-localized transcription factors SOX1 (early neuroectoderm), T/Brachyury, SOX17, and FOXA2 (early mesoendoderm).

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Figure 1B.4.5 Immunohistochemical analysis of formaldehyde-fixed teratoma tissue. In these examples, proteins expressed in ectodermal tissue were localized in tumors developed from human ES cells transplanted into the testis of severe combined immunodeficient mice. Teratomas were grown for a period of 6 to 8 weeks. Serial tissue sections (A,B) were taken through a region of neural differentiation, showing the typical morphology of neuroepithelium organized as neural rosettes (nr; A, stained with HE). Neuroprogenitor cells are thought to reside within the rosette, whereas more mature neural tissues are located around the periphery of these proliferative centers, as indicated by the staining of β-tubulin-III (B), a marker of more mature neural cells. Other types of epithelia identified within the teratoma that lined surfaces and cavities were not neural in nature and possessed a distinctly different morphology (C). These epithelia stained positive for human epidermal keratin (D) but not markers of the neural lineage. Scale bars: (A,B) 150 µm; (C,D) 75 µm.

Isolation of hESC-Derived Teratomas

Gene expression evaluation Teratomas can also be analyzed for gene expression by using an absolute value method such as quantitative RT-PCR for a small set of genes (McDaniel et al., 2007) or a nonquantitative large-scale gene expression method such as hybridization to microarrays (see Basic Protocol 5; UNIT 1B.2). The profiles of expression can then be compared to the published expression patterns of adult tissues in public databases (for example, the NCBI’s GEO database: http://www.ncbi.nlm.nih.gov/geo), and to the expression patterns of the same genes in the hESCs used to generate the teratomas (gene expression profiles for some hESC lines are provided at http://www.stemcellcommunity.org). A teratoma sample should exhibit reduced expression of markers of pluripotency, and elevated expression of markers of mature tissues. Table 1B.4.3 provides a limited list of typical transcript markers and their expected relative levels of expression in undifferentiated hESCs in vitro and differentiated cell types in teratomas in vivo. Additional marker genes that have been used to verify more specific tissues occurring in teratomas include for neuroepithelium (ectoderm)—NEFH (Neurofilament-high molecular weight); TUBB3 (Tubulin, beta 3); bone (mesoderm)—MATN1 (matrilin 1, also known as Cartilage Matrix Protein); muscle (mesoderm)—ACTC (actin, alpha, cardiac muscle, also known as C-Actin) or myosin (myosin light polypeptide 2); pancreas (endoderm)—IPF1 (insulin promoter factor 1, also known as Pancreatic Duodenal homeobox 1); liver (endoderm)—ALB (albumin), or C/EBPα (CCAAT/enhancer binding protein, alpha; Gertow et al., 2006; Schuldiner et al., 2000).

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Table 1B.4.2 List of Commonly Observed Tissues in hESC Teratomas and Some Examples of Primary Antibodies Used to Further Strengthen the Morphological Analysis

Tissue

Antigen

Antibody

Reference

Neurofilament protein (NFP)

DakoCytomation

Gertow et al. (2004)

TuJ1, β-tubulin-III

Promega

Cytokeratins (e.g., CK5)

DakoCytomation

Gertow et al. (2004)

S-100 (Ca -binding protein)

DakoCytomation

Plaia et al. (2006)

Bone sialoprotein (BSP)

Chemicon

Gertow et al. (2004)

Osteocalcin (OCN)

Cambio

Gertow et al. (2004)

Ectodermal derivatives Neuroepithelium Epidermal epithelium Ganglia/Glia

2+

Mesodermal derivatives Bone

2+

Cartilage

S-100 Ca -binding protein

DakoCytomation

Plaia et al. (2006)

Smooth muscle

Smooth muscle actin (SMA)

DakoCytomation 2

Plaia et al. (2006)

Endodermal derivatives Intestine epithelium Lung epithelium

Cytokeratin 8/18a (e.g.Cam5.2) BD Bioscience

Gertow et al. (2004)

a

Cytokeratin 8/18 (e.g.Cam5.2) BD Bioscience b

TTF-1, Thyroid Transc. Fact.-1

Gertow et al. (2004)

DakoCytomation

Gertow et al. (2004)

a Gut and lung epithelia typically stain negative for CK5. b TTF-1 is an endodermal marker present during early thyroid and lung development, later restricted to AT2 (alveolar cells Type 2) and Clara cells

(Van Vliet, 2003).

Table 1B.4.3 Possible Marker Genes to be Used in RT-PCR to Identify Tissues in hESC Teratomasa

HESC

Extraembryonic Endoderm Mesoderm endoderm

Gene name

Symbol

α - fetoprotein

AFP

−

+

+

−

−

Bone morphogenetic protein 4

BMP4

−

−

−

+

−

Cadherin 1 (E-Cadherin)

CDH1

+

+

−

+

+

Cerberus 1

CER1

−

+

+

+

−

Chemokine (C-X-C Motif) Receptor 4

CXCR4

−

−

+

+

−

Disabled 2 homolog

DAB2

−

+

−

+

−

DNA methyltransferase 3B

DNMT3B

+

−

−

−

−

Endometrial bleeding associated factor

EBAF

+

−

−

−

−

Forkhead Box F1

FOXF1

−

−

−

+

−

GATA binding factor 6

GATA6

−

+

−

+

−

Goosecoid

GSC

−

−

+

+

+

Left-right determination factor 1

LEFTY1

+

−

−

−

−

LIM homeobox 1

LHX1

−

+

+

+

−

lin-28 homolog

LIN28

+

−

−

−

−

Ectoderm

continued

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Table 1B.4.3 Possible Marker Genes to be Used in RT-PCR to Identify Tissues in hESC Teratomasa , continued

Extraembryonic Endoderm Mesoderm endoderm

Gene name

Symbol

HESC

Ectoderm

Mesenchyme homeobox 1

MEOX1

−

−

−

+

−

Mix1 homeobox-like 1

MIXL1

−

+

−

+

−

Nanog homeobox

NANOG

+

−

−

−

−

Cadherin 2, N-Cadherin

CDH2

−

−

+

+

−

Nestin

NES

+

−

−

−

+

NK2 transcription factor related, locus 5

NKX2-5

−

−

−

+

−

Nodal homolog

NODAL

±

+

−

+

−

POU domain, class 5, transcription factor 1

POU5F1

+

−

−

−

−

snail homolog 1

SNAI1

−

−

+

+

−

SRY (sex determining region Y)-box 1

SOX1

−

−

−

−

+

SRY (sex determining region Y)-box 17

SOX17

−

+

+

−

−

SRY (sex determining region Y)-box 2

SOX2

+

−

−

−

+

T, brachyury homolog

T

−

−

−

+

−

Undiff. embryonic cell transcription factor 1

UTF1

+

−

−

−

−

Zinc Finger Protein 42

ZFP42

+

−

−

−

−

Zinc Finger Protein of the Cerebellum 1

ZIC1

−

+

−

+

−

Zinc Finger Protein of the Cerebellum 2

ZIC2

−

−

−

+

+

a For sequences and additional information, see Hatta and Takeichi, 1986; Rosner et al., 1990; Wilkinson et al., 1990; Candia et al., 1992; Nieto et al.,

1992; Ang et al., 1993; Monaghan et al., 1993; Sasaki and Hogan, 1993; Zhou et al., 1993; Conlon et al., 1994; Winnier et al., 1995; Candia and Wright, 1996; Nagai et al., 1997; Radice et al., 1997; Varlet et al., 1997; Pevny et al., 1998; Pearce and Evans, 1999; Pesce and Scholer, 2001; Hart et al., 2002; Shook and Keller, 2003; Stemmler et al., 2003; Ellis et al., 2004; Elms et al., 2004; Kunath et al., 2005; Schwartz et al., 2005; Cai, 2006; Li et al., 2006; Liu et al., 2006; Plaia et al., 2006.

BASIC PROTOCOL 4

PARAFORMALDEHYDE FIXATION AND PREPARATION OF TISSUES FOR IMMUNOHISTOCHEMISTRY This protocol describes a method for immersion fixation using paraformaldehyde followed by specimen analysis using immunohistochemistry. In this example the authors have used immunofluorescent detection of specific antigens (see Fig. 1B.4.5). This procedure involves antigen retrieval, but this is not suitable for use with all antibodies. Immunological staining of cryosectioned materials offers an alternative strategy where antigen retrieval does not work, however, the morphology of frozen tissues may not be as well preserved.

Materials

Isolation of hESC-Derived Teratomas

hESC-derived teratomas 4% (w/v) paraformaldehyde (Sigma; 4% (w/v) formaldehyde may also be used) Phosphate-buffered saline (PBS, Sigma) 60%, 70%, 90%, and 95% ethanol Absolute ethanol Histoclear (Sigma) or xylene Paraffin wax and appropriate molds 10 mM citrate buffer (Sigma), pH 6

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Blocking/washing solution: 1% (w/v) bovine serum albumin (BSA, Sigma)/0.2% (v/v) Triton-X-100 (Sigma)/5% (v/v) normal goat serum (Sigma) in PBS Primary antibody Secondary labeled (e.g., FITC-conjugated) antibody Mounting medium: DPX (Sigma) or aqueous mountant (Vectorlabs) Rotary microtome Microscope slides and coverslips (electrostatically charged for improved section adhesion, Sigma) Microwave oven Fluorescence microscope Digital camera and associated imaging software Additional reagents and equipment for fixation, embedding, and sectioning tissues (http://home.primus.com.au/royellis/histo.html or Hofman, 2002) Fix tissues 1. Fix surgically excised tumor tissues by submersion in 4% paraformaldehyde (PFA) in phosphate-buffered saline at 4◦ C for 12 to 24 hr. IMPORTANT NOTE: Optimal tissue preservation can be achieved by perfusion of the deceased animal with 4% PFA prior to surgical removal of the tumor samples followed by submersion fixation as above. See http://home.primus.com.au/royellis/histo.html or Hofman, 2002 for detailed protocols on fixation, embedding, and sectioning.

Embed tissues 2. Process tissues immediately by dehydration through serial ethanol solutions; initiated in 60% to 70% ethanol, progressing through 90% to 95% ethanol, then two or three changes of absolute ethanol before clearing in Histoclear or xylene, followed by embedding in paraffin wax. Section tissues 3. Serially section specimens (5 to 8 µm) using a standard rotary microtome. 4. Mount sections on microscope slides. If necessary, counterstain adjacent serially sectioned samples using hematoxylin and eosin to view morphology.

5. Dewax tissue sections (three washes, each 10 min in Histoclear) and rehydrate through graded ethanol solutions; initiated in absolute ethanol, progressing through 90% to 95% ethanol followed by 60% to 70% ethanol, into water. 6. Perform antigen retrieval by microwaving (800 W) the samples in 10 mM citrate buffer (pH 6) for 2 min and repeat three times. 7. Wash sections in blocking solution for 60 min.

Add primary antibody 8. Incubate sections with primary antibody at the appropriate concentration in blocking solution at 4◦ C overnight. Antibody concentrations and times of incubation may need to be optimized to obtain best results.

9. Wash samples three times in PBS, 2 min each.

Add secondary antibody 10. Incubate samples with secondary conjugated antibody at the appropriate concentration in blocking solution at room temperature for 2 hr.

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11. Wash samples three times in PBS, 2 min each. 12. Coverslip tissue sections using an aqueous mounting medium for fluorescence samples. DPX mountant can be used for chromogenic labels such as diaminobenzidine tetrahydrochloride (DAB).

13. Analyze samples using appropriate microscopy. BASIC PROTOCOL 5

PREPARATION OF TISSUES FOR mRNA EXPRESSION ANALYSIS The following protocol describes how to undertake expression analyses of the teratoma tissues. When the level of gene expression is prioritized over obtaining good tissue morphology, the teratomas can be analyzed using a large-scale gene expression method such as microarrays (UNIT 1B.2).

Materials hESC-derived teratoma, surgically removed Liquid nitrogen (alternatively dry ice/ethanol bath) RNA purification kit (Ambion) Reverse transcriptase Labeled nucleotide Primer composed of oligo(dT) fused to a bacteriophage T7 promoter T7 polymerase Uridine triphosphate (UTP) and Biotin-16-UTP (e.g., Perkin Elmer Life and Analytical Sciences) RNA amplification kit (e.g., the Illumina RNA Amplification kit; Ambion) Amersham Fluorolink streptavidin-Cy3 (GE Healthcare Bio-Sciences) Cryostat 1.5-ml nuclease-free microcentrifuge tubes Microarray chip (e.g., the Refseq 6 BeadChip; Illumina, Inc) Confocal scanner and software (e.g., Illumina BeadArray Reader confocal scanner and software (Illumina BeadArray) 1. Snap-freeze surgically removed hESC-derived teratoma in either liquid nitrogen or dry ice/ethanol bath. 2. Section tissue at 20-µm on a cryostat; collect every tenth section for histology, collect the rest of the sections and place into a 1.5-ml nuclease-free microcentrifuge tube. 3. Isolate total RNA or polyadenylated RNA from the samples with commercial kits that are virtually foolproof. Examples are kits produced, by Ambion and Qiagen. Alternatively, there are numerous acceptable protocols in the literature (Schwartz et al., 2005; Li et al., 2006; Liu et al., 2006). Briefly, the steps performed to isolate RNA are (1) cell lysis and (2) protein-DNA precipitation to separate protein and DNA from the RNA, followed by (3) RNA precipitation, and (4) DNase treatment. DNase treatment is performed since genomic DNA can interfere with primer specificity or cause a false-positive signal in the PCR reaction.

RNA labeling This method is analytical platform-dependent. Isolation of hESC-Derived Teratomas

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4a. cDNA arrays. For cDNA arrays (spotted arrays), convert RNA to single-stranded cDNA using reverse transcriptase and a labeled nucleotide, which is incorporated into the cDNA during the reaction. This labeled cDNA is then used directly to hybridize to the array. Current Protocols in Stem Cell Biology

4b. Oligonucleotide arrays. Convert RNA to cDNA using a primer composed of oligo(dT) fused to a bacteriophage T7 promoter. Use T7 polymerase to drive transcription from the incorporated promoter to produce RNA from each cDNA. Include a labeled nucleotide during this reaction, to provide the ability to detect a fluorescent signal bound to array elements. This second technique (described above) requires linear amplification of the RNA several hundred to >1000-fold, and is used for oligonucleotide arrays, such as Illumina’s 50-mer arrays, Agilent’s 70-mer arrays, and Affymetrix’s 20- to 25-mer arrays. The amplification methods are beneficial when starting materials are limited. For the Illumina BeadArray system, labeling is done by incorporating biotin-16-UTP (Perkin Elmer Life and Analytical Sciences), present at a ratio of 1:1, with unlabeled UTP.

5. Hybridize target to arrays. Hybridization is usually an overnight incubation, and the arrays are washed the next day (see manufacturer’s instructions; UNIT 1B.2). The incubation conditions, buffers, and washing steps vary by array manufacturers. All manufacturers sell the buffers either as part of their array kits or as separate packages, or they are made in the laboratory following the recipes provided by the manufacturers. For the Illumina BeadArray system, labeled, amplified material (700 ng per array) is hybridized to the Illumina Refseq 6 BeadChip according to the manufacturer’s instructions (Illumina). Arrays are washed, and then stained with Amersham fluorolink streptavidinCy3 (GE Healthcare Bio-Sciences) according to methods provided by the manufacturer.

Perform data extraction, processing, and normalization 6. Use a confocal scanning fluorescence microscope to scan hybridized arrays and capture the signals. The pixels corresponding to array elements are identified by image analysis software, which then extracts hybridization signals, generating a table of values for each gene. For most systems this is a fully automated process that takes no more than a few minutes. The extracted pixels are condensed to a single value for each transcript, often incorporating background subtraction or other data processing algorithms. Array signals are often normalized within an experiment to even out differences in overall intensity or other technical variations. For the Illumina BeadArray system hybridized arrays are scanned using an Illumina BeadArray Reader confocal scanner, and array data processing and analysis are performed using Illumina BeadStudio software. Differential expression of individual genes between groups is calculated by the t-test.

Data analysis The data that is generated can be analyzed using software as simple as spreadsheets to enterprise-wide array database and analysis systems. Currently certain standards in microarray analysis are emerging. This topic is too broad to survey adequately here but is covered in a recent review (Allison et al., 2006; UNIT 1B.2). Most biologists want to start with a hit list of up or down regulated genes in their different experimental conditions. This approach is straightforward and freely available software packages for this type of question are available and have become accepted as standard for certain applications. The authors suggest Significance Analysis for Microarrays (SAM) as an excellent tool to obtain lists of genes that are up or down regulated within a given dataset. The advantage of SAM is that it provides the experimenter not only with significance levels for results but also with sample size assessment—estimates of false discovery rate (FDR), false negative rate (FNR), type I error, and power for different sample sizes and other features. For further reading, see e.g., the SAM Web site: http://www-stat.stanford.edu/∼tibs/ SAM/. Current Protocols in Stem Cell Biology

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COMMENTARY Background Information By definition, a teratoma is a benign tumor of germ cell origin composed of somatic tissues derived from all three germ layers (ectoderm, mesoderm, and endoderm; Stevens, 1967). Xenografting of karyotypically normal hESC to immunodeficient mice generates a similar benign growth (Thomson et al., 1998) and has been generally referred to as the formation of hESC teratoma. Immunodeficient mutant mice as recipients To avoid rejection of the xenograft, treatment of the recipient animal with immunosuppressants could be used (Grinnemo et al., 2006). More commonly however, is to use mutant mice with immunodeficiencies. Immune-deficient SCID (C.B-17-Prkdcscid ) mice were explored in 1983 by Bosma et al. (1983). Similar to nude mice, the SCID mice do have NK cells, macrophages, and neutrophils. However, a DNA repair defect due to a mutant gene responsible for recombination of (VDJ) segments of the T- and B-cell receptors results in B and T cell deficiency. These SCID mice may have residual B and T cell activity which is why even more efficient modified severe combined immunodeficient (SCID) mice were developed. Mice with the SCID mutation were crossed with NK-deficient beige mice (Roder and Duwe, 1979) that have a defect in lysosomal trafficking reducing also NK-cell function, and phagocytosis (Mosier et al., 1993). The SCID/beige crosses N7) (C.B-Igh-1b GbmsTac-Prkdcscid -Lystbg therefore have a double-mutation causing impaired lymphoid development and reduced natural killer cell activity, and these mice are commonly used as host for the growth of hESC-derived teratomas (Thomson et al., 1998; Gertow et al., 2004). Mice with only the SCID mutation have also been used to induce teratomas (Reubinoff et al., 2000; Stojkovic et al., 2004a). Other immune-deficient mouse models include the NOD/ShiJic-SCID with γ cnull (NOD/SCID/γ c null ) mouse that has multiple immunological dysfunctions (Ito et al., 2002). This is a cross with nonobese diabetic (NOD) mice that have a defect in antigen presentation and T-cell function (Makino et al., 1980).

Isolation of hESC-Derived Teratomas

Pros and cons of the hESC-teratoma model system With in vitro culture today it is not possible to achieve the complexity needed for the full

cellular differentiation into tissues similar to in vivo formation of teratomas. Although new approaches for cell growth in vitro, such as three-dimensional scaffolds (Levenberg et al., 2003; Hayman et al., 2004), are a step forward, many of the environmental cues that modulate cellular differentiation are absent in in vitro models. Differentiation following xenografting of hESCs is a better option; however, it is still limited when compared to injection of mESCs into a blastocyst. In hESC-derived teratomas, growth is three-dimensional, but correct embryo folding and movements of cells for normal development does not occur, resulting in spherical and haphazard tissues. The differentiation time frame and the volume of the tissue that can be allowed to develop are limited. This is due to limitations on what the host can supply with regard to vascularization and space, but also to what is ethically reasonable for the specific animal model used. More advanced structural elements can be found with longer times, but there is also increased necrosis due to crowding. The condition of the injected cells obviously affects the outcome; this is the reason it is very important for these studies to be performed with well-defined cells. The reproducibility of the hESC injection from the same cell line is high, but hESC lines appear to differ in their developmental capacity by gross morphology (Heins et al., 2004; Przyborski, 2005) and differentiation (Mikkola et al., 2006). Conventional HE histology is rarely sufficient to accurately define structures in the teratomas. Marker studies, using immunocytochemistry, are particularly useful to determine the maturation level and identity of cells or tissues otherwise not recognizable. In general, more mature tissues require larger sets of markers for their identification and determination of accurate cell components and position. However, it must be noted that biomarkers cannot be used to infer functionality.

Critical Parameters and Troubleshooting Testicular or kidney capsule rupture during injection of hESCs If the volume of cells injected is too large this will cause the injected content to be pushed out of the injection hole (see Fig.1B.4.1; testis), or even make the capsule burst. Therefore it is critical to keep the volume as low as possible.

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Mortality during anesthesia and surgery Hypothermia. Mice have a high surface area relative to body mass, consequently they are highly susceptible to hypothermia, and they lose body temperature very easily. This can be avoided by providing heat (heat blanket or lamp) and by keeping the anesthesia as short as possible. Inadequate depth or lethal dose of anesthesia. The effective dose of anesthesia needed may vary. The depth of anesthesia must be carefully followed for each individual. Bleeding. As little as 0.5 ml loss of blood may cause cardiovascular failure (Flecknell, 1993). Replacement of fluids with sterile saline must be considered. Mortality 1 to 5 days after surgery Common causes of post-surgical mortality may be infections or fluid loss. The risk for infections can be reduced by aseptic handling during surgery and the use of sterile equipment. Fluids may be replaced. Lack of teratoma growth Generally, lack of teratoma growth indicates (1) that the injected cells lacked developmental potential (as an endogenous trait or from detrimental handling during preparations), or (2) that the implantation procedure failed technically. Possible causes of technical failure include (1) differentiated hESC, metabolic stress, dead cells; (2) cells stayed inside the injection needle, too few cells injected; and (3) too much fluid (and too few cells) injected leading to leakage of cells and poor engraftment (particularly in the kidney capsule). Host contribution to tissues in the teratoma-like growth To verify hESC origin of the observed tissues, it is advisable to perform immunohistochemistry with the human-specific antibodies; e.g., anti-E-cadherin (Zymed Laboratory) or anti–human nuclei (Chemicon).

Anticipated Results In general, a 100% rate of hESC teratoma formation should be expected. In the authors’ experience, reproducibility from repeated injections of the same hESC line is also close to 100%, i.e., similar composition of tissues are formed. hESC lines differ, however, in their in vivo pluripotency and there is no known correlation between pluripotency detected in vitro and in vivo. In addition, the growth and differentiation of hESCs appears to be influenced by the graft site (Cooke et al., 2006).

The following tissues are most often identified in hESC-derived teratomas: 1. Primitive epithelium, and neuroepithelium, and retina-like pigmented epithelium; 2. Structures consisting of a simple columnar epithelium, crypts with proliferative cells, mucus-producing goblet cells, and smooth muscle layers; 3. Loose mesenchymal tissue; 4. Cartilage and bone in various stages of development; and 5. Smooth muscle, glands of various types, and immature blood vessels.

Less commonly encountered tissues include: 1. Renal tissue such as tubules and glomeruli 2. Liver 3. Skin cells Several structures produced within teratomas derived from hESCs are highly organized and consist of ordered arrangements of different tissue types that in many ways recapitulate organogenesis within the embryo. For example, the authors and others have reported organized structures that have the appearance of kidney, containing renal corpuscles, associated tubules, and associated vascular supply; gastrointestinal tract consisting of a simple columnar epithelium, supporting mucosa, smooth muscle layers, and neural ganglia; skin including dermal and epidermal layers, complete with stratum granulosum, keratinized cells, and hair follicles; and respiratory airway composed of pseudostratified ciliated epithelium, smooth muscle, nerves, and supporting cartilage (Figs. 1B.4.3, 1B.4.4, and 1B.4.5; Gertow et al., 2004; Stojkovic et al., 2004b). There are also other examples of tissue types found in isolation within the body of the teratoma that may be identified using standard histological methods, including skeletal muscle, neural ganglia, pigmented cells, glands, primitive epithelium, and neuroepithelium (Figs. 1B.4.3, 1B.4.4, and 1B.4.5 Gertow et al., 2004; Stojkovic et al., 2004b). Accordingly, the engraftment of hESCs into an appropriate host can result, in part, in the differentiation of human tissues that consist of cells in a recognized arrangement that resemble structures within the developing embryo and adult.

Time Considerations hESC teratoma formation is a long-term experiment that needs to be planned well ahead.

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The following are some important practical issues to be considered: hESCs It is pivotal to have access to the hESCs in logarithmic growth phase for the transplantation. For this, it is recommended (using your standard culture protocol) to passage the hESCs 2 to 3 days before the injection. Avoid transporting the cells unless good culture conditions can be maintained during the transport. It is the authors’ experience that hESC transported for >1 hr at room temperature do not engraft as readily. Mice If the recipient mice need to be transported to your facility, they must arrive 1 to 2 weeks before the start of the experiment for acclimatization. Time for HESC-derived teratoma tissue to develop Teratoma growth is dependent on the cell concentration of the inoculum. When 104 to 105 cells are implanted the hESC-teratomas can generally be harvested after 6 to 10 weeks, or may be grown further up to 12 weeks without adding discomfort to the animal. This timing can however differ slightly between hESC lines. Teratomas from higher doses, >106 cells, can generally be harvested within 6 weeks.

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Key References Hofman, F. 2002. See above. Detailed knowledge on histology and collections of protocols used in histology. Van Zutphen, L.F.M., Baumans, V., Beynes, A.C. 2001. Principles of Laboratory Animal Science; Revised edition. Amsterdam, Netherlands. This book covers the main theoretical aspects of laboratory animal science. Waynforth and Flecknell, 1999. See above. This book covers standard surgical procedures

Internet Resources http://iacuc.cwru.edu/policy/nihpolicies/ surguide.htm To learn more about animal experimentation, particularly rodent surgery, the authors recommend NIH Guidelines for Rodent Surgery. http://home.primus.com.au/royellis/histo.html Detailed knowledge on histology and collections of protocols used in histology. http://www.ncbi.nlm.nih.gov/geo Published expression patterns of adult tissues. http://www.stemcellcommunity.org Provides expression profiles of the same genes in the HESCs. http://www-stat.stanford.edu/∼tibs/SAM/ Further reading on Significance Analysis for Microarrays (SAM) used to obtain lists of genes that are up or down regulated within a given microarray dataset.

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Tandem Affinity Purification of Protein Complexes in Mouse Embryonic Stem Cells Using In Vivo Biotinylation

UNIT 1B.5

Jianlong Wang,1 Alan B. Cantor,1 and Stuart H. Orkin1,2 1

Children’s Hospital and the Dana Farber Cancer Institute, Harvard Medical School, Harvard Stem Cell Institute, Boston, Massachusetts 2 The Howard Hughes Medical Institute, Boston, Massachusetts

ABSTRACT Streptavidin affinity purification of protein complexes, in combination with in vivo biotinylation of critical transcription factors, has contributed to the analysis of the pluripotent state in mouse embryonic stem (ES) cells and made it possible to construct a proteinprotein interaction network.This has facilitated discovery of novel pluripotency factors and a better understanding of stem cell pluripotency. Here we describe detailed procedures for an in vivo biotinylation system setup in mouse ES cells, and affinity purification of multi-protein complexes using in vivo biotinylation. In addition, we present a protocol employing SDS-PAGE fractionation to reduce sample complexity prior to submission for mass spectrometry (MS) protein identification. Curr. Protoc. Stem Cell Biol. 11:1B.5.1C 2009 by John Wiley & Sons, Inc. 1B.5.17.  Keywords: tandem affinity purification r in vivo biotinylation r protein-protein interaction r embryonic stem cells

INTRODUCTION To maintain essential cellular functions, a large number of individual proteins must assemble into an array of multi-protein complexes of different structure and composition and act in a coordinated fashion. In order to understand stem cell pluripotency and other complex biological phenomena, it is essential to analyze and characterize protein complexes and elucidate intricate protein-protein interaction networks. There are a number of different methods for purifying proteins and other large molecules of interest from complex mixtures, e.g., crude extracts. One of these is affinity purification, which exploits specific binding interactions between molecules. The first step in affinity purification is to obtain a ligand of the target molecule immobilized on a solid support material and incubate the crude sample with the immobilized ligand, thereby allowing the target molecule in the sample to bind to the ligand attached to the support material. Non-bound sample components are then washed away, and the target molecule, together with its associated proteins, is eluted by altering the buffer conditions in such a way that the binding weakens or ceases to occur. A common strategy for affinity purification is to use two different affinity tags in a tandem purification. The FLAG peptides DYKDDDDK and MDYKDDDDK are widely used affinity tags (Chubet and Brizzard, 1996) that can be placed at either the amino-terminus or the carboxy-terminus, or in association with other tags such as the biotinylation peptide tag (see Background Information). The protocols in this unit are based on our earlier studies using in vivo biotinylation to perform affinity purification of pluripotency factors and construct a pluripotency network in mouse ES cells (Wang et al., 2006). Additional protocols covering protein-protein and protein-DNA interactions have been presented in Kim et al. (2009). The general strategy is summarized in Figure 1B.5.1 and Figure 1B.5.2. This unit begins with a method

Current Protocols in Stem Cell Biology 1B.5.1-1B.5.17 Published online October 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b05s11 C 2009 John Wiley & Sons, Inc. Copyright 

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Figure 1B.5.1 Establishment of a biotinylation system in J1 ESCs. A stable ESC line expressing the bacterial BirA enzyme was first established by transfection with a BirA-expressing plasmid bearing the neomycin resistance (neor ) gene and G418 selection. A second plasmid containing cDNA encoding a transcription factor (TF) of interest with an N-terminal Flag-biotin dual tag (FLBIO) and a puromycin resistance (puror ) gene was introduced, and cells were selected with puromycin. The resulting stable line is resistant to both G418 and puromycin, and expresses FLAG-tagged, biotinylated TF (FLBIO TF) that can be immunoprecipitated by anti-FLAG and streptavidin antibodies/beads.

Tandem Affinity Purification of Protein Complexes in mESC

Figure 1B.5.2 A summary of the procedure for tandem affinity purification of multiprotein complexes in mouse ESCs. Following establishment of BirA-only and BirA+FLBIO TF-expressing ES cell lines, immunoprecipitation is performed using anti-FLAG M2 agarose (FLAG-IP). The bound material is eluted with FLAG peptide and further purified by streptavidin affinity capture. The purified protein complexes are fractionated on SDS-PAGE, and subjected to LC-MS/MS to identify components of the protein complexes.

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to establish an in vivo biotinylation system in mouse ES cells (see Basic Protocol 1), followed by a detailed protocol to perform tandem affinity purification of the biotinylated protein together with its associated protein complexes (see Basic Protocol 2). Finally, a detailed protocol for fractionation of purified protein complexes (to increase sample purity and reduce sample complexity) for downstream mass spectrometry analysis is presented (see Support Protocol). NOTE: All ES cell cultures should be maintained at 37◦ C in a humidified atmosphere of 5% CO2 in air. NOTE: For tandem affinity purification, all reagents and solutions should be kept on ice unless otherwise specified.

ESTABLISHMENT OF ES CELL LINES EXPRESSING BirA AND SUB-ENDOGENOUS BIOTINYLATED PROTEINS

BASIC PROTOCOL 1

Two cell lines are generated in this protocol: ES cells expressing BirA only are established first to be used as control cells for background signals during affinity purification of protein complexes. This cell line also serves as the recipient cells for subsequent introduction of genes dually tagged with Flag and biotin tags (FLBIO). By first establishing BirA-only ES cell lines using G418 selection, future introduction of different FLBIO-tagged genes (using puromycin selection) and thus the establishment of multiple FLBIO-tagged cell lines in the presence of same amount of BirA expression is made possible. Nuclear extracts are prepared simultaneously from BirA-expressing cells with and without tagged genes of interest, with simultaneous affinity purification (Basic Protocol 2).

Materials J1 ES cells (ATCC, cat. no. SCRC-1010) ES medium (see recipe) IEF medium (see recipe) 0.05% (w/v) trypsin (Mediatech, cat. no. 25-052-CI) 0.25% (w/v) trypsin (Mediatech, cat. no. 25-053-CI) Phosphate-buffered saline (PBS; Sigma, cat. no. D8537) pEF1αBirAV5-neo plasmid (see Fig. 1B.5.1; available from the author upon request) TE buffer (see recipe) 300 μg/ml G418 (from 300 mg/ml stock; see recipe) 2× freezing medium (see recipe) RIPA buffer (Boston BioProducts, cat. no. BP-115) anti-V5-HRP (Invitrogen, cat. no. 46-0708) pEF1αFlagbiotin (FLBIO)-puro plasmid (see Fig. 1B.5.1; available from the author upon request) Puromycin (see recipe) Streptavidin-HRP (Amersham, cat. no. RPN1231) 6-well, 24-well, 48-well, and 10-cm IEF plates (see recipe) 15-ml conical tubes (Corning, cat. no. 430791) 0.4-cm gap cuvette for electroporator (Bio-Rad, cat. no. 165-2008) Gene Pulsor II (electroporation; Bio-Rad) 37◦ C, 5% CO2 incubator U-bottom 96-well plate 200-μl pipettor Multi-channel pipettor (e.g., 12-channel pipettor) Parafilm Gelatin-coated cell culture plates (see recipe)

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Establish cultures 1. Using standard cell culture procedures, thaw J1 ES cells (or ES cells of your choice) and expand to ∼70% confluence in a well of a 6-well IEF plate containing 5 ml ES medium per well. It can take a few days to a week to expand the cells depending on the starting cell number.

2. One the day prior to electroporation, prepare a 10-cm IEF plate with 10 ml IEF medium (for use in step 10 to plate the transformed cells). Mouse embryonic feeder cells (MEFs) are used in the culture of mouse ES cells. They both provide a substrate for the ES cells to grow on and secrete many factors necessary for ES cells to maintain pluripotency. Feeders are MEFs that have been mitotically inactivated by treatment with mitomycin C or by γ -irradiation (Conner, 2000; UNIT 1C.3). A unique quadresistant DR4 feeder cell line can be purchased from Open Biosystems (cat. no. MES3948), which has been mitotically inactivated by treatment with mitomycin C. Alternatively, we also isolated primary embryonic fibroblast from DR4 mouse embryos, expanded them and inactivated them by γ -irradiation. These will be referred to as irradiated embryonic feeders (IEF) in this unit.

Prepare cells for electroporation 3. To harvest cells by trypsinization, aspirate ES medium and rinse the cells once with 0.05% trypsin. Then, add a sufficient amount of 0.25% trypsin to cover the ES cells and incubate 3 to 5 min at 37◦ C. The ES cells should become detached from the vessels; if not, increase incubation time.

4. Collect cells by adding 3 vol ES cell medium to neutralize the trypsin and pipetting up and down to mix. Transfer to 15-ml conical centrifuge tubes. 5. Centrifuge 5 min at 200 × g, 4◦ C. 6. Wash harvested cells twice, each time with 5 to 10 ml PBS. 7. Count cells and then resuspend ES cells at 1.3 × 107 /ml in PBS.

Electroporate cells to produce BirA cells 8. Add 20 to 30 μg of pEF1αBirAV5-neo DNA in no more than 50 μl TE buffer to make final total 0.75 ml of cell suspension (or ∼107 cells) for each electroporation. Note that linearization of the plasmid prior to electroporation is not necessary.

9. To perform electroporation for J1 ES cells, use a 0.4-cm gap cuvette, 25 μF, 450 V, with the time constant for each electroporation reading around 0.6 to 0.8 msec. Incubate 5 min on ice. Different ES cell lines may require different electroporation conditions.

Plate the cells and select transformed cells 10. Mix the electroporated cells with enough ES medium to bring the final cell suspension volume to 10 ml. Remove the medium from the 10-cm IEF plate prepared a day before and add the electroporated cell suspension. Rock the plates gently to mix and then incubate at 37◦ C, 5% CO2 . 11. On the second day (24 hr after initial plating), add 300 μg/ml (final) G418 drug directly to the ES medium on the cells. Rock the plate gently to mix drug completely with the medium and return to the incubator. Tandem Affinity Purification of Protein Complexes in mESC

12. Feed the cells with fresh G418 drug and ES medium each day for the next 7 to 9 days. Swirl the plate gently to resuspend and remove the dead cells and debris by aspiration. Replace with 10 ml freshly made drug-containing ES medium mix to the plate.

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After 3 or 4 days of this treatment, most of the ES cells will appear dead. By day 5 or 6, clones should start appearing and should be ready for picking by day 8 or 9.

Pick clones 13. A day prior to picking, prepare a 48-well IEF plate with 500 μl ES medium per well (for use in step 17) and a U-bottom 96-well plate containing 50 μl of 0.05% trypsin per well (for use in step 15). It is wise to set up the 48-well plate and 96-well plate with matching wells for convenient multi-channel pipettor transfer.

14. On picking day, wash the 10-cm plate with 10 ml PBS. Replace the wash with 5 ml fresh PBS to protect the cells from drying out during picking. 15. Pick individual colonies with a 200-μl pipettor set at 10 μl and transfer each colony into a well of the U-bottom 96-well plate containing 50 μl of 0.05% trypsin. 16. After picking 24 to 48 clones or 30 min (whichever is first), incubate the 96-well plate 10 min at 37◦ C. 17. Using a multi-channel pipettor, transfer 150 μl ES medium from the 48-well IEF plate to the 96-well trypsin plate containing the colonies. Pipet up and down to mix and then transfer the entire cell suspension back to the 48-well feeder plate. Culture the cells in the 48-well plate overnight. 18. Replace old medium with 500 μl fresh ES medium without G418 every day. G418 (300 μg/ml) may be added but is not necessary during the first few days.

Freeze cells 19. When the majority (>70%) of the wells are ready (i.e., cells reach near or over 70% confluency), wash with 500 μl PBS, add 35 μl of 0.05% trypsin, and incubate 10 min at 37◦ C. Meanwhile, prepare a 96-well plate containing 65 μl cold 2× freezing medium. It is wise to set up the 96-well plate to match the 48-well plate (from step 17) for convenient multi-channel pipettor transfer.

20. Using a 12-channel pipettor, add 65 μl ES medium to each well of the 48-well plate, mix by pipetting up and down to neutralize the trypsin, and transfer 65 μl to the U-bottom 96-well plate containing 65 μl cold 2× freezing medium. Wrap the plate in Parafilm and store at −80◦ C for use in step 24. 21. Add 200 μl ES medium to the remaining cells in the IEF plate, and return the plate to the incubator. Replace old medium with fresh ES medium daily for 3 to 4 days.

Analyze clones 22. When the medium in most wells is yellow or cells are near confluence, aspirate medium and add 200 μl RIPA buffer to each well to make total lysate. Carry out SDS-PAGE and a standard western blot analysis with the lysate (20 μg) using antiV5-HRP (1:5000 dilution is recommended). Since BirA is V5-tagged, you should expect to see a band near 35 Kda in the positive clones (see Fig. 1B.5.3A). It is also recommended to make total lysate from the parental ES cells (e.g., J1) for use in western blotting as a negative control.

Expand positive clones 23. Prepare a 24-well IEF plate with 1 ml ES medium per well a day prior to step 24. 24. Thaw the positive clones (as determined by western blotting in step 22) by adding 100 μl of warm ES medium to the frozen wells (from step 20). Mix and transfer

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Figure 1B.5.3 Examples of western blot analyses. (A) A western blot with anti-V5-HRP to detect BirAV5 expression. (B) A western blot using native antibody against the TF of interest to detect both endogenous and biotinylated TF proteins. The sub-endogenous level of the biotinylated transcription factor (FLBIO TF) and the expression level of the endogenous protein (end TF) are indicated. NS denotes nonspecific signals. Panel B is reprinted from Kim et al. (2009) with permission of Nature Protocols.

the thawed cell suspension to 24-well IEF plate prepared a day before (see step 23), repeat until all the remaining cells are thawed and transferred. Return to incubator and culture overnight. 25. On the second day, replace old medium with fresh ES medium containing 300 μg/ml G418. Let clones grow in ES cell selection medium (ES medium with 300 μg/ml G418) until they are 70% confluent (this takes ∼1 week). 26. Freeze down 60% of the cells as frozen stocks and grow the remaining 40% in 10-cm IEF plates until 70% confluent for use in step 27. Alternatively, all the cells can be frozen and stored at −80◦ C (short-term) or in liquid nitrogen (long-term) for later use.

Establish FLBIO(gene) cell line 27. Repeat steps 2 to 26 to establish ES cell lines expressing BirA and biotinylated proteins of interest at sub-endogenous levels. 28. Electroporate BirA-containing cells (from step 26) with pEF1αFlagbio-tagged plasmid containing a specific gene of interest and use ES medium containing 300 μg/ml G418 and 1 to 2 μg/ml puromycin to select for cells expressing both pEF1αBirAV5neo and pEF1αFLBIO(gene)-puro plasmids (see Fig. 1B.5.1). Bacterial BirA will catalyze biotinylation of the FLBIO(gene), which has a biotinylation site.

29. Perform western analysis using a streptavidin-HRP conjugate (to detect the biotinylated protein) and a native antibody against the protein of interest if available (to detect both the biotinylated and endogenous versions of the protein; see an example in Fig. 1B.5.3B); thaw and expand the positive clones to make frozen cell stocks. Milk must not be used during incubation with streptavidin-HRP antibody (see Commentary).

30. Adapt the two cell lines established thus far to grow on gelatin-coated culture vessels without feeders by serially passaging them in complete ES medium. Freeze down gelatin-adapted cell stocks. Tandem Affinity Purification of Protein Complexes in mESC

We found that J1 ES cells could be efficiently adapted to grow on gelatin-coated culture vessels in the presence of LIF after three passages. Alternatively, it is advantageous to start with previously established, feeder-independent ES cells (e.g., E14 line) in the beginning.

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TANDEM AFFINITY PURIFICATION OF PROTEIN COMPLEXES Individual proteins often participate in the formation of a variety of different protein complexes that are a cornerstone of many biological processes. This protocol details large-scale expansion of mouse ES cells and nuclear extract preparation from the bulk culture for use in tandem affinity purification. This purification may require 20 to 40 culture dishes (15-cm) of ES cells grown to near 90% confluence, which corresponds to the yield of 50 to 100 mg of nuclear extract. It is advisable to adapt the two cell lines established in Basic Protocol 1 to grow in a feeder-independent condition (on gelatin) to minimize the cross-contamination by feeder cells and cost of the experiment.

BASIC PROTOCOL 2

Materials Gelatin-coated ES cell culture dishes (see recipe) BirA-only and BirA+ Flagbiotagging ES cells (established in Basic Protocol 1) ES medium (see recipe) 0.05% (w/v) trypsin (Mediatech, cat. no. 25-052-CI) 0.25% (w/v) trypsin (Mediatech, cat. no. 25-053-CI) Phosphate-buffered saline (PBS; Sigma, cat. no. D8537) Nuclear extract buffer A (see recipe) Protease inhibitor cocktail (Sigma Mammalian Protease Inhibitor cocktail) Trypan blue (Invitrogen, cat. no. 15250-061) Nuclear extract buffer B (see recipe) Bradford assay: Protein concentration Bio-Rad Dye kit (Bio-Rad, cat. no. 500-0006) IP350 buffer with different NP40 concentrations (see recipes) Protein G–agarose (Roche, cat. no. 11-243-233001) FLAG M2-agarose beads (Sigma, cat. no. A2220-5ML) FLAG peptide (Sigma, F-3290) Streptavidin-agarose beads (Invitrogen, cat. no. 15942-050) 2× SDS sample buffer (see recipe) 250-ml conical plastic bottles (Corning, cat. no. 430776) Centrifuge with a JS 4.2 rotor or equivalent 50-ml conical tubes (Corning, cat. no. 430829) Glass Dounce homogenizer (40-ml size) with type B pestle (Wheaton, cat. no. 432-1273) Drawn-out glass Pasteur pipet Glass Dounce homogenizer (15-ml size) with type B pestle (Wheaton, cat. no. 432-1272) NALGENE high-speed centrifuge tube (cat. no. 3114-0050) Rotating wheel (Scientific Equipment Products, cat no. 60448) 15-ml conical tubes (Corning, cat. no. 430791) 1.5- and 2-ml screw-cap tube Additional reagents and equipment for counting cells using a hemacytometer (Phelan, 2006), determining cell viability using trypan blue staining (Strober, 1997), and determining protein concentration using the Bradford assay (Siu et al., 2008) Expand the cell lines 1. Culture ES cells as follows: a. Begin with a 10-cm gelatin-coated dish containing 1 × 106 cells/cm2 in 10 ml ES medium for both control BirA and biotinylated protein (from step 30 in Basic Protocol 1). Incubate dishes.

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b. When cells reach 90% confluence, trypsinize and split at a 1:1 ratio into a 15-cm gelatin-coated dish. Incubate dishes. c. When these cells reach 90% confluence, trypsinize and split the 15-cm dishes at a 1:4 or 1:5 ratio into fresh 15-cm gelatin-coated dishes. Incubate dishes. d. Expand cell culture until you have 20 to 40 15-cm gelatin-coated dishes of cells for each cell line. Drug selection is needed only during initial culture and can be omitted during the scaleup culture to reduce costs. Optimal starting material has not been determined; however, you want to have enough cells to make 50 to 100 mg of nuclear extracts (see below).

2. Grow cells until they reach 80% to 90% confluency with constant medium renewal. Ideally, you want to renew the medium on a daily basis; however, you can renew the medium every other day before cells reach 70% confluence to reduce costs. When medium turns very yellow within a day after medium renewal, cells have likely reached near 80% to 90% confluence, and you should go directly to step 3.

Harvest the cells 3. To trypsinize cells, rinse each dish with 5 ml of 0.05% trypsin, then add 7 ml of 0.25% trypsin per dish and incubate the plate 5 min at 37◦ C. Process the dishes ten dishes at a time.

4. Add 10 ml of fresh ES medium to the cells to neutralize the trypsin, resuspend well, and transfer cells to a 250-ml conical plastic bottle. Rinse the dishes with an additional 10 ml of ES medium, collect the residual cells, and combine with the cells in the bottle. RPMI or low glucose DMEM with 10% FBS can be used instead of ES cell culture medium to neutralize trypsin. LIF is not required in the neutralization medium.

5. Centrifuge the cells in 250-ml conical plastic bottles 15 min at 2400 × g (using a JS 4.2 rotor or equivalent), 4◦ C. 6. Carefully decant the supernatant, resuspend the cell pellet in 50 ml of ice-cold PBS, and transfer to 50-ml conical tubes. Count cell numbers using a hemacytometer (Phelan, 2006). Pool pellets if multiple tubes are used for harvesting the same samples (BirA versus BirA+FLBIO TF).

7. Centrifuge the 50-ml conical tubes 10 min at 2400 × g, 4◦ C in a JS 4.2 rotor. 8. Remove the supernatant carefully. Estimate the packed cell volume (PCV). Resuspend in ∼5 PCV volume of ice-cold PBS. Centrifuge again 10 min at 2400 × g, 4◦ C in a JS 4.2 rotor.

Isolate the nuclei 9. Remove the supernatant, and rapidly resuspend in ∼5 PCV of ice-cold nuclear extract buffer A with freshly added DTT (1 mM final), PMSF (0.2 mM final), and 1:1000 protease inhibitor cocktail. 10. Centrifuge 5 min at 2400 × g, 4◦ C in a JS 4.2 rotor.

Tandem Affinity Purification of Protein Complexes in mESC

11. Aspirate the supernatant carefully. Add ∼3 PCV of ice-cold nuclear extract buffer A and additives described in step 9 (except use 1:100 protease inhibitor cocktail). Incubate on ice for 10 min to swell cells. 12. Transfer by pouring into a glass Dounce homogenizer (40-ml size) with type B pestle prechilled on ice and prerinsed with nuclear extract buffer A. Homogenize up and down 10 times, slowly.

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Check for cell lysis by examination of small aliquot with trypan blue stain (Strober, 1997) under microscope (>80% of cells should be lysed).

13. Centrifuge 15 min at 4300 × g, 4◦ C using a JS 4.2 rotor.

Extract the nuclei 14. Carefully remove and discard the supernatant using a drawn-out glass Pasteur pipet. Add to the nuclei pellet 3 ml/1 × 109 starting number of cells of ice-cold nuclear extract buffer B containing freshly added DTT (1 mM final), PMSF (0.2 mM final), and protease inhibitor cocktail (1:1000). 15. Dislodge the pellet and transfer to a glass Dounce homogenizer (15-ml size) with type B pestle, prechilled and rinsed with nuclear extract buffer B. Homogenize up and down 10 times slowly to resuspend the pellet. 16. Transfer the homogenate to a NALGENE centrifuge tube, prerinsed with nuclear extract buffer B and rotate for 30 min on a rotating wheel in the cold room (4◦ C). 17. Centrifuge 30 min at 25,000 × g, 4◦ C using a JA-25.50 rotor to remove insoluble material. Meanwhile, prepare for the Bradford assay according to manufacturer’s instructions. 18. Carefully transfer the supernatant (contains nuclear extract) to 50-ml conical tubes.

Determine protein concentration 19. Determine the protein concentration of an aliquot of each nuclear extract (NE) using the Bradford assay (Siu et al., 2008; e.g., the Bio-Rad Dye kit). Preclear nuclear extract 20. Use equal amounts (∼100 mg) of NE from the BirA (control) sample and BirA + biotinylated protein sample. Add an appropriate amount of cold IP350 buffer (0.3% or 0.5% v/v NP-40) to each sample so that the final NE concentration of each sample is ∼2 mg/ml containing ∼0.2% NP-40. Addition of DTT/PMSF/protease inhibitor cocktail to the IP350 buffer is preferred but not necessary.

21. Preclear supernatant with Protein G–agarose (100 μl of 50% slurry per 10 mg protein) for 1 to 2 hr at 4◦ C with continuous mixing. 22. Centrifuge samples 5 min at 300 × g, 4◦ C. Transfer precleared supernatant to new 50-ml conical tubes.

Affinity purify FLAG-tagged proteins 23. Equilibrate FLAG M2 agarose beads (100 μl of 50% slurry per 10 mg protein) in two 50-ml tubes containing 15 ml cold IP350/0.3% NP-40 buffer. Rotate in cold room for 5 min and centrifuge 4 min at 300 × g, 4◦ C. Aspirate. Repeat this once. 24. Carefully transfer the precleared nuclear extract prepared in step 21 to preequilibrated FLAG M2 resin from step 23. Divide into more 50-ml tubes if necessary to ensure complete mixing during immunoprecipitation. Place on end-over-end rotating wheel at 4◦ C overnight. 25. Centrifuge tubes from overnight incubation 4 min at 300 × g, 4◦ C. Remove supernatant (nonbound material). Make sure no obvious protein precipitate (whitish clumps) is formed after overnight incubation. Embryonic and Extraembryonic Stem Cells

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26. Wash FLAG-agarose beads four times, each time with 20 ml of ice-cold IP350/0.3% NP-40 buffer. Place on rotating wheel at 4◦ C for 15 min per wash. Centrifuge 4 min at 300 × g, 4◦ C between washes. 27. After the final wash, remove most of the supernatant. Use IP350/0.3% NP-40 buffer to transfer beads to a 15-ml tube. Pool beads, if multiple tubes were used for the incubation and washes from previous steps. Centrifuge 4 min at 300 × g, 4◦ C to pellet beads. Remove the supernatant carefully. 28. Elute beads four times, each time with 10 ml of IP350 with 0.1 mg/ml FLAG peptide. For each elution, place sample tubes on rotating wheel for 1 to 1.5 hr at 4◦ C. Centrifuge 4 min at 300 × g, 4◦ C, and carefully transfer the supernatant (contains eluted proteins) to a new 50-ml conical tube. Pool the eluates for each sample for a total 40 ml of each sample. Discard beads.

Affinity purify biotin-labeled protein 29. Prepare streptavidin agarose, as described for FLAG M2 agarose in step 23. 30. Add the FLAG-eluate (40 ml each) from step 28 into equilibrated streptavidin– agarose and place on end-over-end rotating wheel at 4◦ C overnight. 31. Centrifuge tubes from overnight incubation 4 min at 300 × g, 4◦ C. Remove the supernatant (nonbound material). Make sure no obvious protein precipitate (whitish clumps) is formed after overnight incubation.

32. Wash streptavidin-agarose beads four times, each time with 20 ml of ice-cold IP350/0.3% NP-40 buffer. Place on rotating wheel for 15 min at 4◦ C per wash. Centrifuge as described in step 31. 33. After the final wash, remove most of the supernatant. Use a cut-off pipet tip and remaining IP350 buffer to transfer beads into a 1.5-ml screw-cap tube. Pool beads, if multiple tubes were used for the incubation and washes. Wash out original tubes and pipet tips with IP350 buffer and pool with sample. Centrifuge using tabletop microcentrifuge 2 min at 300 × g, 4◦ C to pellet beads. 34. Remove as much supernatant as possible from the beads using a drawn-out Pasteur pipet. Add 500 μl of 2× SDS sample buffer. Vortex gently and heat at 95◦ to 100◦ C for 5 min and vortex again. Allow to cool to room temperature. 35. Centrifuge 1 min at maximum speed, room temperature, to repellet beads. Carefully transfer supernatant to a new 1.5-ml screw-cap tube. Add 500 μl of 1× SDS sample buffer to residual beads and vortex gently. Recentrifuge 1 min at maximum speed, room temperature. Pool the sample with the first eluate in a 1.5-ml screw-cap tube. 36. Repeat washing of beads one more time with 400 μl of 1× SDS sample buffer and combine with samples (should now have 1.4 ml total). Repeated washing of the beads after boiling (and combining the supernatants) improves the yield of proteins.

37. Centrifuge 2 min at maximum speed, room temperature (to pellet any residual agarose beads that were carried over). Carefully transfer sample into new 2-ml screw-cap tubes. Tandem Affinity Purification of Protein Complexes in mESC

Carryover of residual agarose beads will block the Centricon filter (see Support Protocol). Samples can be frozen and stored at −80◦ C for future use.

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SDS-PAGE FRACTIONATION OF PROTEIN COMPLEXES This protocol details the concentration of the protein eluates and fractionation of the purified protein complexes on a SDS-PAGE gel to reduce the complexity of samples for downstream mass spectrometry analysis.

SUPPORT PROTOCOL

Materials Affinity-purified complexes with biotinylated FLAG-tagged protein (Basic Protocol 2) 30% Acrylamide/Bis solution (37.5:1) (Bio-Rad, cat. no. 161-0158) Colloidal Coomassie stain (Invitrogen, cat. no. 46-7015 and 46-7016) HPLC-grade water (American Bioanalytical) YM-10 Centricon (10,000 MWCO; Amicon Bioseparations, cat. no. 4205) 37◦ C water bath Avanti J25 centrifuge using a JA-25.50 fixed-angle rotor 1.5-ml microcentrifuge tubes Bio-Rad Protean II xi basic unit with casting stand (Bio-Rad, cat. no. 165-1834) Scalpel or razor blade Additional reagents and equipment for preparing a large denaturing polyacrylamide gel (Gallagher, 2006) 1. Transfer all eluate (from step 37, Basic Protocol 2) into the chamber of a YM-10 Centricon (10,000 MWCO) device. If just thawed from the freezer, warm samples in a 37◦ C water bath for 5 min to dissolve SDS. Undissolved SDS will block the Centricon filter.

2. Centrifuge in the Avanti J25 centrifuge using a JA-25.50 fixed-angle rotor ∼2 to 3 hr (until as much filtrate runs through the chamber as possible) at 5000 × g, 25◦ C . After centrifugation there should be ∼100 μl of concentrated material. This centrifugation step needs to be performed at room temperature to avoid precipitation of the SDS in the samples.

3. Remove and discard filtrate chamber first. Attach collection vial and invert quickly. Re-centrifuge inverted chamber (with collection vial) 2 min at 800 × g, 25◦ C. Transfer to a fresh 1.5-ml microcentrifuge tube. 4. Prepare a large denaturing polyacrylamide gel: Use Bio-Rad Protean II xi basic unit with casting stand. Prepare gel with fresh APS and TEMED. Pour lower running gel (10%) first and seal with ethanol or butanol for polymerization. Next, pour upper stacking gel (4%) with comb positioned in an angle, and then reposition the comb horizontally to avoid air bubbles in the comb wells. Pull out the bottom gel blocker carefully before use (Gallagher, 2006). 5. Load samples from step 3 and run the gel for ∼1 hr at 120V in stacking gel and run samples till the bromphenol blue dye reaches ∼2.5 cm into the separating (lower) gel (it takes ∼1 hr under 120V). Fill in empty wells with the same volume of 1× protein sample buffer and a similar salt concentration to prevent “smiling” effect from sample wells. Also, protein ladder can be loaded on left and right sides of the gel; the ladders help to cut out gel slices with an expected size range of proteins for mass spectrometry (see step 8).

6. Disassemble gel apparatus and stain gel overnight with Colloidal Coomassie stain.

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7. Destain gel with multiple rinses of HPLC-grade water. Destain until background is very clear (can take 1 to 2 days) on rocker platform at room temperature. 8. Cut out the whole lane and separate into four to eight slices for mass spectrometry analysis. Using a scalpel or razor blade, cut into ∼1-mm cubes and transfer to clear 1.5-ml microcentrifuge tubes. Add a small amount of HPLC-grade water so that gel slices do not dry out. Do not spin tubes to avoid samples leaching out of gels. Cut-out gel slices can be stored at 4◦ C (or −20◦ C for prolonged storage).

9. Send samples to a MS facility of your choice to perform liquid chromatography coupled with tandem MS (LC-MS/MS) for protein identification.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

ES medium Dulbecco’s modified Eagle’s medium (DMEM) 15% (v/v) fetal bovine serum (FBS; Hyclone, cat. no. SH30071.03) 0.1 mM 2-mercaptoethanol 2 mM L-glutamine 0.1 mM non-essential amino acid 1% (v/v) nucleoside mix (100× stock, Sigma) 1000 U/ml recombinant leukemia inhibitory factor (LIF; Chemicon) 50 U/ml penicillin/50 μg/ml streptomycin (Invitrogen, cat. no. 15070-063) Store up to 1 month at 4◦ C Each lot of fetal bovine serum needs to be prescreened for the ability to support optimal ES cell growth.

Freezing medium, 2× 20% (v/v) dimethyl sulfoxide (DMSO) 80% (v/v) fetal bovine serum (FBS; Hyclone, cat. no. SH30071.03) Store up to 1 month at 4◦ C G418 Also known as geneticin, G418 (Invitrogen, cat. no., 11811) is a broad-spectrum antibiotic that will select mammalian cells expressing the neomycin protein (encoded by the neomycin gene). Make a 300 mg/ml stock solution in PBS. Store up to 6 months at −20◦ C.

Gelatin, 0.1% Dissolve 5 g of gelatin (Bacto; Difco, cat. no., 0143-15-1) in 500 ml distilled water and autoclave (1% stock). Store the solution at room temperature indefinitely. Before use, dilute 1:10 (to make 0.1% working solution) with sterile dH2 O and filter through a 0.45-μm filter apparatus.

Gelatin-coated tissue culture plates Tandem Affinity Purification of Protein Complexes in mESC

Add a sufficient amount of 0.1% gelatin (see recipe) to each well of the culture plates, let stand for 20 min to 1 hr. Aspirate the gelatin solution and air dry the plates. Store the gelatin-coated plates at room temperature indefinitely.

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IEF medium 86% (v/v) DMEM (high glucose) 10% fetal bovine serum (FBS; heat inactivated) 2% (v/v) penicillin/streptomycin 5000 U penicillin/5000 μg/ml streptomycin; Invitrogen, cat. no. 15070-063 1% of 200 mM L-glutamine 1% of 100 mM sodium pyruvate (Invitrogen, cat. no. 11360-070) Store up to 1 month at 4◦ C IMPORTANT NOTE: It is extremely important that the sodium pyruvate is fresh. Using an expired lot will decrease your yields by 50% or more.

IEF plates Normally, 3 × 106 IEF cells are frozen down as 1× stock. When thawed, these cells are used to seed one full gelatin-coated tissue culture plate (i.e., 1 × 96-well plate—3 × 104 cells/well, 1 × 48-well plate—6.25 × 104 cells/well, 1 × 24-well plate—1.2 × 105 cells/well, 1 × 12-well plate—2.4 × 105 cells/well, 1 × 6-well plate—4.8 × 105 cells/well, or 1 × 10-cm dish—3 × 106 cells/dish).

IP350 buffer (0.3% NP-40) 350 mM NaCl 20 mM Tris·Cl, pH 7.5 0.3% (v/v) NP-40 1 mM disodium EDTA 10% (v/v) glycerol Store up to several months at 4◦ C Just before use, add fresh 1 mM DTT, 0.2 mM PMSF, protease inhibitor cocktail (1:1000; Sigma Mammalian Protease Inhibitor cocktail) IP350 buffer (0.5% NP-40) 350 mM NaCl 20 mM Tris·Cl, pH 7.5 0.5% (v/v) Nonidet P40 (NP-40) 1 mM disodium EDTA 10% (v/v) glycerol Store up to several months at 4◦ C Just before use, add fresh 1 mM DTT, 0.2 mM PMSF, protease inhibitor cocktail (1:1000; Sigma Mammalian Protease Inhibitor cocktail) Nuclear extract buffer A 20 mM HEPES 10 mM KCl 1 mM disodium EDTA 0.1 mM Na3 VO4 0.2% (v/v) Nonidet P40 (NP-40) 10% (v/v) glycerol Store up to several months at 4◦ C Just before use add fresh 1 mM DTT, 1 mM PMSF, and protease inhibitor cocktail (1:1000; Sigma Mammalian Protease Inhibitor cocktail)

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Nuclear extract buffer B 20 mM HEPES 10 mM KCl 1 mM disodium EDTA 0.1 mM Na3 VO4 350 mM NaCl 20% (v/v) glycerol Store up to several months at 4◦ C Just before use, add fresh 1 mM DTT, 1 mM PMSF, protease inhibitor cocktail (1:1000) NaCl concentration can vary from 150 mM to 500 mM, which needs to be determined empirically.

Puromycin Make 1 mg/ml puromycin (Sigma, cat. no., P8833) in PBS. Filter sterilize using a 0.22-μm filter, divide into aliquots, and store up to 6 months at −20◦ C. Freeze/thaw stock no more than 5 or 6 times.

SDS sample buffer, 2× 1% (v/v) glycerol 3% (w/v) SDS 0.5 M Tris·Cl, pH 6.8 0.004% (w/v) bromphenol blue Store up to 6 months at −20◦ C TE buffer 10 mM Tris·Cl, pH 7.5 1 mM disodium EDTA Store up to 12 months at room temperature (25◦ C) COMMENTARY Background Information

Tandem Affinity Purification of Protein Complexes in mESC

Mammalian protein complexes have been studied by combining protein affinity purification with mass spectrometry (MS) and bioinformatics. Particularly useful among the affinity-based purification methodologies is the biotin-avidin system. Protein biotinylation is a powerful technique for molecular biology and biomedical applications due to the high affinity and specificity of the biotin-avidin interaction (Cronan, 1990; Beckett et al., 1999). For the purpose of protein purification, biotinylation affords a number of advantages. First, the strong affinity of biotin for avidin allows purification of the biotinylated protein under high stringency conditions, thus minimizing background binding that may be observed with a different affinity tag or by using native antibodies. A second advantage is that naturally occurring biotinylated proteins are rare, making cross-reactions less likely to happen. Thirdly, with this approach, there is no need to generate protein specific-antibodies,

eliminating the risk of such antibodies crossreacting with cellular proteins other than those intended. Biotinylation can occur either by the cell’s endogenous protein-biotin ligases, or through the coexpression of an exogenous biotin ligase (e.g., the bacterial BirA enzyme), as described in this protocol. Biotinylation of a tagged transcription factor mediated by an exogenous biotin ligase BirA has been demonstrated by others to maintain the factor’s protein interactions, DNA binding properties in vivo, and subnuclear distribution (de Boer et al., 2003). Recently, the authors of this unit assessed the usefulness of in vivo biotinylation of transcription factors in mouse embryonic stem (ES) cells. First, we established an approach for the single-step and tandem purification of transcription factor complexes based on specific in vivo biotinylation mediated by BirA (Wang et al., 2006). Second, we demonstrated the feasibility of in vivo biotinylation for mapping global/chromosomal targets of many

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different transcription factors (Kim et al., 2008). A notable point is that the same cells expressing a biotin-tagged version of a given transcription factor can be utilized for the construction of both protein-protein and proteinDNA interaction networks (unpub observ.). Although the authors of this unit performed all of these studies in mouse ES cells, the approaches should be adaptable for working with other types of cells.

Critical Parameters In Basic Protocol 1, gelatin adaptation to make ES cells feeder-independent is important for the following two reasons: (1) it eliminates contamination by feeder cells in subsequent purification; (2) it greatly reduces the experimental cost incurred by the large-scale culture of ES cells required for affinity purification of protein complexes. Be aware that not all ES cells are favorable for gelatin adaptation and feeder-independent growth, so selection of ES cell lines to start with that can be gelatin adapted (e.g., J1 ES cells) or grow without feeders (e.g., E14 cells) is advantageous. To screen for the positive clones expressing biotinylated protein, it is critical not to add milk during streptavidin-HRP antibody incubation, since the milk may contain biotinrelated species that can interfere with the streptavidin antibody. Ideally, western analysis with the native antibody should be performed to detect relative expression level of the biotinylated protein versus endogenous protein, and only the clones with sub-endogenous expression levels should be selected for affinity purification (see an example in Fig. 1B.5.3). The selection of sub-endogenous expression levels of tagged protein ensures minimal interference with endogenous protein complexes by the tagged protein and thus allows for affinity purification of the bona fide interacting partners. However, the native antibody is not always available for your protein of interest; in this case, several clones with medium- or low-level expression of biotinylated proteins should be used for affinity purification. During affinity purification of protein complexes (Basic Protocol 2), a sufficient amount of the starting nuclear extract (NE) is important to obtain enough final material for MS analysis. A good starting point is 50 to 100 mg of NE protein. In addition, the salt concentration in nuclear extraction buffer B is critical for solubility and integrity of protein complexes. A salt concentration that is too low can result in protein precipitation and aggregation during overnight incubation of nuclear extracts with

affinity agarose beads; a salt concentration that is too high may disrupt weak protein-protein interactions. The salt concentration used in affinity purification can vary from 150 mM to 500 mM; therefore, the optimal salt concentration for your protein of interest has to been determined empirically. In the Support Protocol, due to the high sensitivity of mass spectrometry, it is critical to use high-quality HPLC-grade water to stain and destain the SDS-PAGE gel. Always wear gloves when handling gels, and keep gels in a dish covered with Saran wrap to avoid contamination. In addition, an experienced, reliable MS facility is critical to ensure the success from this expensive and time-consuming experiment. Like all the biological experiments, the same affinity purification procedure should be repeated twice or three times to ensure reproducibility, and to increase the likelihood of identifying bona fide protein-protein interactions.

Troubleshooting See Table 1B.5.1 for troubleshooting tips.

Anticipated Results The authors of this unit typically screen 24 to 48 G418-resistant clones for expression of BirAV5 using anti-V-HRP antibody when using the in vivo biotinylation setup. More than 50% of these clones will show one level or another of BirA expression. Given that BirA is an active biotin ligase, the authors have determined that the Flagbiotin (FLBIO)– tagged gene product was biotinylated efficiently, mediated by low-, medium-, and highBirA-expressing cells. The authors screen similar numbers of BirA and Flagbiotag clones; one-quarter to one-third should be positive for the FLBIO tag, as determined by means of streptavidin-HRP antibody. Tandem affinity purification usually results in the observation of minimal background binding signals in BirA control cells and few endogenously biotinylated proteins in BirA+Flagbiotag cells. Detailed information on the BirA control background signals and the endogenously biotinylated proteins is available in de Boer et al. (2003) and Wang et al. (2006). Ranging from a few to >10 candidates, the total number of potential interaction partners of the tagged protein varies from one individual protein to another. The candidate proteins are usually of high confidence; however, it is necessary to obtain further validation

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Table 1B.5.1 Troubleshooting Guide for Tandem Affinity Purification of Complexes from ES Cells

Problem

Possible reasons

Solution

Weak or no positive signals for the Milk contains high levels of biotin, Western blot analysis with streptavidin-HRP biotinylated proteins (Basic which block the interaction should be done without milk Protocol 1, step 22) between streptavidin and the biotinylated protein Low yield of nuclear extract from both control BirA and BirA+Flagbiotin-tagged samples (Basic Protocol 2, step 19)

High nonspecific binding present in both BirA only and BirA+Flagbiotin-tagged samples

No true positive clones

Screen more clones

Cell lysis is not complete

Make sure cell lysis is complete at step 12

Low cell numbers

Optimize cell culture and increase the starting material

The NE concentration is too high

Dilute/adjust the NE concentration to ∼2 mg/ml

% of NP-40 in IP350 is too low

Increase % of NP-40 in IP350

Protein precipitates after overnight The salt concentration is not incubation (Basic Protocol 2, steps optimal 25 and 31) The final volume does not go down to 100 μl (Support Protocol 1, step 2)

There are still residual resins in the Briefly centrifuge the eluate, recover the eluate, which clogs the Centricon supernatant carefully, and extend the device centrifuge time at step 2 SDS precipitates

by reverse tagging followed by affinity purification and/or co-immunoprecipitation.

Time Considerations

Tandem Affinity Purification of Protein Complexes in mESC

Adjust NaCl concentration in nuclear extraction buffer B. It can vary from 150 mM to 500 mM. The optimal salt concentration has to be determined empirically.

In vivo biotinylation system setup The most time-consuming procedure described in this unit is the in vivo biotinylation system setup. This phase must be planned meticulously and carried out in a timely manner. Generally, it takes ∼50 days to set up the system as described in this unit. The setup process can potentially be shortened to ∼30 days by establishing both the BirA-only and the BirA+Flagbiotag ES cell lines simultaneously. If this is done, it will be necessary to select for the BirA-only line with G418 only after PEF1αBirAV5-neo electroporation, and to select for the BirA+Flagbiotag line with G418 and puromycin together, after co-electroporation with the PEF1αBirAV5neo plasmid and the plasmid containing the PEF1αFLBIO-tagged gene of interest.

Warm up the eluates before Centricon filtration

Tandem affinity purification This depends on the scale of culture and starting ES cell numbers. From a 10-cm culture dish, it takes over a week (∼10 days) to expand to twenty 15-cm dishes. It takes another full day to make nuclear extracts. In general, we prepare nuclear extract, determine protein concentration, and set up Flag M2-agarose or streptavidin-agarose incubation all on the same day, to avoid prolonged storage and/or freezing/thaw cycles that will potentially disrupt multiprotein complexes. An extra day is required for a second affinity purification with streptavidin agarose. Therefore, allow ∼2 weeks for executing the full protocol of tandem affinity purification. SDS-PAGE fractionation of protein complexes for MS spectrometry The concentration of the protein eluates using Centricon takes a few hours, and the SDSPAGE fractionation takes another few hours. Once the samples are fractionated on gels, the

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staining is done overnight, and the de-staining takes from one to a few days. The gel slices can be cut out and stored at 4◦ C for the short term before being submitted for mass spectrometry analysis.

de Boer, E., Rodriguez, P., Bonte, E., Krijgsveld, J., Katsantoni, E., Heck, A., Grosveld, F., and Strouboulis, J. 2003. Efficient biotinylation and single-step purification of tagged transcription factors in mammalian cells and transgenic mice. Proc. Natl. Acad. Sci. U.S.A. 100:7480-7485.

Acknowledgements

Gallagher, S. 2006. One-dimensional SDS gel electrophoresis of proteins. Curr. Protoc. Molec. Biol. 75:10.2A.1-10.2A.38.

This work is supported by Seed Grant from the Harvard Stem Cell Institute Cell Reprogramming Program to J.W. A.B.C. is supported by NIH Grant R01 HL075705. S.H.O. is an Investigator of Howard Hughes Medical Institute.

Literature Cited Beckett, D., Kovaleva, E., and Schatz, P.J. 1999. A minimal peptide substrate in biotin holoenzyme synthetase-catalyzed biotinylation. Protein Sci. 8:921-929. Chubet, R.G. and Brizzard, B.L. 1996. Vectors for expression and secretion of FLAG epitopetagged proteins in mammalian cells. Biotechniques 20:136-141.

Kim, J., Cantor, A.B., Orkin, S.H., and Wang, J. 2009. Use of in vivo biotinylation to study protein-protein and protein-DNA interactions in mouse embryonic stem cells. Nat. Protoc. 4:506517. Kim, J., Chu, J., Shen, X., Wang, J., and Orkin, S.H. 2008. An extended transcriptional network for pluripotency of embryonic stem cells. Cell 132:1049-1061. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Siu, F.K.Y., Lee, L.T.O., and Chow, B.K.C. 2008. Southwestern blotting in investigating transcriptional regulation. Nat. Protoc. 3:51-58.

Conner, D.A. 2000. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Molec. Biol. 51:23.2.1-23.2.7.

Strober, W. 1997. Trypan blue exclusion test of cell viability. Curr. Protoc. Immunol. 21:A.3B.1A.3B.2.

Cronan, J.E., Jr. 1990. Biotination of proteins in vivo. A post-translational modification to label, purify, and study proteins. J. Biol. Chem. 265:10327-10333.

Wang, J., Rao, S., Chu, J., Shen, X., Levasseur, D.N., Theunissen, T.W., and Orkin, S.H. 2006. A protein interaction network for pluripotency of embryonic stem cells. Nature 444:364-368.

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Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

UNIT 1B.6

Jennifer A. Erwin1,2,3 and Jeannie T. Lee1,2,3 1

Howard Hughes Medical Institute, Boston, Massachusetts Department of Molecular Biology, Massachusetts General Hospital, Boston, Massachusetts 3 Department of Genetics, Harvard Medical School, Boston, Massachusetts 2

ABSTRACT This unit describes a method of performing ßuorescent in situ hybridization (FISH) of XIST and Cot-1 RNA in human pluripotent stem cells (hPSC) to characterize the epigenetic status of X-chromosome inactivation (XCI). hPSC laboratories commonly practice karyotypic analysis to monitor genetic stability; however, epigenetic stability is often overlooked. Several laboratories have recently shown that markers of XCI can be used as one effective screen to monitor the epigenetic status of hPSCs. Human embryonic stem cells (HESC) fall into three classes of XCI states: upregulating XIST upon differentiation, always expressing XIST in the undifferentiated and differentiated states, and never expressing XIST in the undifferentiated and differentiated states. Failure to express XIST represents an especially concerning state in hESC, as this state does not occur in healthy female cells but is often seen in malignancies. Herein, methods of carrying out XIST RNA and Cot-1 RNA FISH are described. FISH analysis of XIST RNA, unlike general expression analysis such as RT-PCR, allows for the classiÞcation of XCI on a single-cell level, enabling a quantitative determination of the degree of epigenetic change across the population. The complementary Cot-1 analysis measures the extent of repeat element expression throughout the nucleus and therefore enables determination, at a cytological level, of the extent to which the X chromosome is silent. Because the different steps of XCI are some of the Þrst epigenetic changes to take place in differentiating hESC, analysis of the XCI state provides a Þrst indication of an hESC C 2010 by John culture’s overall health. Curr. Protoc. Stem Cell Biol. 12:1B.6.1-1B.6.11.  Wiley & Sons, Inc. Keywords: human embryonic stem cells r epigenetics r X-chromosome inactivation r XIST

INTRODUCTION Adaptation of hPSC to prolonged culture can lead to a variety of genetic and epigenetic changes. Because XCI is one of the Þrst epigenetic changes to take place in differentiating hESC, characterization of XCI status can serve as a marker of the epigenetic stability within a cell line or culture. Herein, methods to characterize XCI status by assaying for XIST RNA expression in undifferentiated and differentiated cells (XIST RNA FISH) while also assaying for active transcription off of the X chromosome by Cot-1 RNA FISH are described. hPSCs vary in their epigenetic status with respect to XCI, with different XCI statuses reported between different laboratories, different cultures, or even different cells within the same culture. This unit begins with the isolation, slide preparation, and Þxation of hPSCs (see Basic Protocol 1), then follows with the in situ staining for RNA transcripts by RNA FISH (see Basic Protocol 2), and ends with the staining of X-chromosomes by DNA FISH

Current Protocols in Stem Cell Biology 1B.6.1-1B.6.11 Published online February 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b06s12 C 2010 John Wiley & Sons, Inc. Copyright 

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(see Alternate Protocol). The Support Protocols describes labeling XIST (see Support Protocol 1) and Cot-1 DNA (see Support Protocol 2) and preparation of the hybridization probe mixture (see Support Protocol 3). NOTE: Equipment and reagents should be sterile and aseptic technique should be used. NOTE: The following procedures involving live cells are performed in a Class II biological hazard ßow hood or a laminar-ßow hood. NOTE: For the purpose of RNA-FISH, all reagents and labware should be RNAse-free. BASIC PROTOCOL 1

ISOLATION, SLIDE PREPARATION, AND FIXATION OF HUMAN PLURIPOTENT STEM CELLS This protocol is used to generate hPSC slides for use in RNA or DNA FISH. Colonies are broken up enzymatically to single cells, cells are spun onto slides with a cytospin centrifuge, and cells are then permeabilized and Þxed. Slides can be stored up to 3 weeks for RNA FISH and for 12 months for DNA FISH at 4◦ C.

Materials hPSCs cultures (starting with at least 1 × 105 cells per hybridization spot) Phosphate buffered saline without CaCl2 and without MgCl2 (CMF-PBS), room temperature and ice cold 0.05% (w/v) trypsin/EDTA MEF medium (see recipe) CSK-T solution (see recipe), ice cold 4% paraformaldehyde in 1× CMF-PBS, pH 7.2 (4% PFA) 70% ethanol, ice cold 37◦ C incubator 5-ml serological pipets 15-ml conical tubes (Falcon) Cytospin centrifuge Positively charged slides (e.g., Fisherbrand Superfrost/Plus) Coplin jars Isolate hPSC 1. Aspirate medium from 1 well of hPSCs cultured in a 6-well organ culture dish, and gently rinse the plate with 2 ml CMF-PBS. 2. After aspirating CMF-PBS, cover plate with 0.5 ml 0.05% trypsin/EDTA and incubate 5 min at 37◦ C. Cells can be harvested from a variety of culture conditions, including growth on feeders or Matrigel. An 80% conßuent well will yield enough cells for 15 to 20 slides.

3. Inactivate the trypsin by adding 4 ml MEF medium. Gently pipet up and down with a 5-ml serological pipet to break up the colonies. Over trypsinization or rough pipetting compromises the slide quality.

Count cells 4. Transfer cells and medium to a 15-ml conical tube. Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

5. Remove 20 μl of cell/medium to count cells with Trypan blue to test cell viability and cell number (UNIT 1C.3). 6. Centrifuge remainder of cell suspension 5 min at 200 × g, room temperature. 7. Resuspend cells in CMF-PBS at a concentration of 1 × 106 cells/ml.

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Prepare slides 8. Assemble cytospin apparatus with positively charged slides. 9. Transfer 100 μl of cell suspension/CMF-PBS solution to cytofunnel. 10. Centrifuge in the cyotspin for 5 min at 1000 rpm at room temperature. 11. Disassemble the cytospin apparatus and allow slide to dry for 5 min. The slide must be dry to attach the cells to the slide.

12. Immerse slides in ice-cold CMF-PBS in a Coplin jar for 5 min. 13. Transfer slides to ice-cold CSK-T in a Coplin jar and incubate 3 min. 14. Transfer slides to room temperature 4% PFA in a Coplin jar and incubate 10 min. 15. Transfer slides to ice-cold 70% ethanol in a Coplin jar. The slides can now be stored in 70% ethanol for 3 weeks at 4◦ C (RNA FISH) or for several months at 4◦ C (DNA FISH). For long-term storage, the slides can be dehydrated in ethanol (see Basic Protocol 2) and stored at −80◦ C.

XIST DNA PROBE PREPARATION BY NICK TRANSLATION This section describes the labeling of XIST. Silencing of the X chromosome is reßected by exclusion of Cot-1 RNA from the inactive X-chromosome and XIST territory. During XCI, XIST RNA coats the inactive X chromosome. Thus, presence of XIST RNA also reßects X-chromosome inactivation. XIST DNA is labeled using the nick translation method, which uses DNase I to introduce randomly distributed nicks into DNA. DNA polymerase I then uses the nicked 3 -OH as a primer to synthesize complementary DNA, while incorporating Cy3-labeled dUTP.

SUPPORT PROTOCOL 1

Materials XIST exon1 DNA (GenBank U80460: 61251–69449) Nick translation kit (Roche, cat. no. 10976776001) containing: dATP dGTP dCTP 10× buffer Enzyme mix Cy3-dUTP (Amersham, cat. no. PA53022) PCR thermal cycler 1. Add 2 μg of probe DNA (XIST exon1) to the following components in the nick translation kit:

3 μl dATP (0.4 mM) 3 μl dGTP (0.4 mM) 3 μl dCTP (0.4 mM) 1.2 μl Cy3-dUTP (1 mM) 2 μl 10× buffer 2 μl enzyme mix 20 μl total volume 2. Incubate reaction 2 hr at 15◦ C. 3. Inactivate the reaction by incubating 15 min at 65◦ C in a thermal cycler.

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4. Store probe (up to 1 year) at –20o C before continuing on to Support Protocol 3. This reaction is good for 25 hybridizations. SUPPORT PROTOCOL 2

COT-1 PROBE LABELING BY RANDOM PRIMING Cot-1 DNA is labeled by random priming using the Stratagene Prime-It Fluor kit. Cot-1 DNA is denatured and primed with random 9-mer probes, and the Klenow fragment then extends the primers into probe fragments while incorporating ßuor-12-dUTP. This protocol produces enough label for 25 hybridizations.

Materials Cot-1 DNA (Invitrogen, cat. no. 15279-011) Prime-It Fluor with FITC-dUTP kit (Stratagene, cat. no. 300380) containing: Random 9-mers 5× nucleotide buffer Fluor-12-dUTP Klenow Stop buffer PCR thermal cycler 1. Add 2 μg of Cot-1 DNA to 10 μl of random 9-mers from the Prime-It Fluor with FITC-dUTP kit and 26 μl of DEPC-treated H2 O. 2. Incubate 5 min at 95◦ C in a thermal cycler. 3. Brießy spin 1 min at 16,000 × g, room temperature, and place on ice for 5 min. 4. Add the following components from the kit:

9.2 μl 5× nucleotide buffer 0.8 μl ßuor-12-dUTP 2 μl Klenow 50 μl total volume 5. Incubate 30 min at 37◦ C. 6. Add 2 μl stop buffer. 7. Immediately proceed to Support Protocol 3 or store for 1 year at –20◦ C. SUPPORT PROTOCOL 3

DNA PRECIPITATION OF LABELED PROBES FOR ANALYSIS OF X-CHROMOSOME INACTIVATION For analysis of XCI, the reaction mixes containing Cy3-dUTP-labeled XIST DNA probe and the ßuor-12-dUTP-labeled Cot-1 DNA probe are combined. The labeling reaction is precipitated with herring sperm DNA and resuspended in hybridization buffer.

Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

Materials Cy3-dUTP-labeled XIST DNA probe reaction mix (see Support Protocol 1) Fluor-12-dUTP-labeled Cot-1 DNA reaction mix (see Support Protocol 2) Herring sperm DNA (Promega, cat. no. D1811) 3 M sodium acetate, pH 5.0 70% and 100% ethanol Hybridization buffer (see recipe) 1. Once both probes are labeled separately, mix the total products from the labeling reactions in Support Protocols 1 and 2 (20 μl of XIST probe and 52 μl of Cot-1) together and add 40 μg (4 μl) herring sperm DNA.

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2. Precipitate with sodium acetate and ethanol by adding 7.6 μl of 3 M sodium acetate and 152 μl of 100% ethanol. Freeze 30 min at −20◦ C. 3. Centrifuge 30 min at 16,100 × g, 4◦ C. DNA should be precipitated into a pellet at the bottom of the tube.

4. Remove the supernatant, making sure not to disturb the pellet. 5. Wash DNA by adding 200 μl of 70% ethanol (at 4◦ C). 6. Centrifuge at 16.1 × g for 5 min at 4◦ C. 7. Remove supernatant and air dry the pellet. 8. Resuspend the probe in 200 μl of hybridization buffer to a Þnal concentration of 20 ng (calculated from input DNA) of each probe DNA per microliter. Store the probe up to 1 year at −20◦ C protected from light.

X-CHROMOSOME INACTIVATION DETECTED BY RNA FLUORESCENT IN SITU HYBRIDIZATION

BASIC PROTOCOL 2

This protocol describes the hybridization, washing, and detection of the XIST and Cot-1 RNA signals. The probe must be pre-annealed before adding to the slide and the slides should be completely dehydrated through an ethanol dehydration series. Also, the overnight hybridization on the slides must occur in a humidiÞed chamber. Once mounted, the signal is detectable for several weeks.

Materials XIST/Cot-1-labeled probe (see Support Protocols 1 through 3) Fixed slides in 70% ethanol (see Basic Protocol 1) 80%, 90%, and 100% ethanol, ice cold 2× SSC/50% (v/v) formamide (see recipe for 20× SSC) 2× SSC (see recipe for 20× SSC) Vectashield with DAPI (Vector) Clear nail polish 42◦ C and 80◦ C heating blocks or water baths HumidiÞed chamber at 37◦ C (e.g., pipet tip container with water in the bottom) Glass coverslips Coplin jars 45◦ C incubator with agitator Upright epißuorescence microscope with 60× oil immersion lens or confocal microscope equipped with Þlters (compatible for imaging with DAPI, Cy3, Cy5, and FITC) Pre-anneal XIST probe 1. From Support Protocol 3, transfer 8 μl of XIST/Cot-1 probe (20 ng/μl) for each hybridization spot. 2. Denature labeled probe 10 min at 80◦ C. 3. Incubate probe 5 to 60 min at 42◦ C for, while dehydrating the slides (steps 4 to 8).

Dehydrate slides 4. Transfer slides to ice-cold 80% ethanol and incubate 5 min. 5. Transfer slides to ice-cold 90% ethanol and incubate 5 min. 6. Transfer slides to ice-cold 100% ethanol and incubate 5 min.

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7. Air dry slides. 8. Place slides in pre-warmed (42◦ C) humidiÞed chamber.

Hybridize slides 9. Add 8 μl of pre-annealed probe (at 42◦ C) to the spot of cells on the slide. Carefully cover the probe with a coverslip, avoiding bubbles. 10. Incubate 12 hr at 42◦ C.

Wash slides 11. Pre-warm three Coplin jars with 2× SSC/50% formamide and three coplin jars with 2× SSC 20 min at 45◦ C. 12. Remove the coverslip. Wash slides three times in 2× SSC/50% formamide 5 min each time at 45◦ C with light agitation. 13. Wash slides three times in 2× SSC 5 min each time at 45◦ C with light agitation.

Detect signal 14. Apply 10 μl Vectashield with DAPI mounting medium to each cell spot. Seal coverslip with clear nail polish. 15. Visualize slides with an upright epißuorescence microscope with a 60× oil immersion lens or with a confocal microscope equipped with Þlters that are compatible for imaging with DAPI (400 nm), Cy3 (570 nm), Cy5 (660 nm), and FITC (505 nm). Store the slides at 4◦ C in the dark. Signals are stable for a minimum of 6 months. Refer to Silva et al. (2008) for examples of staining patterns. ALTERNATE PROTOCOL

DNA FLUORESCENT IN SITU HYBRIDIZATION RNA FISH procedures used to assay XCI are often accompanied by DNA FISH to conÞrm that the analysis is carried out in euploid cells with two X-chromosomes. Only one inactive X should occur in diploid cells with two X chromosomes. This protocol describes the detection of X chromosomes in interphase nuclei by DNA FISH with X-chromosome paint as a probe. This protocol can follow the RNA FISH protocol, in which case the RNA FISH signal is Þxed with PFA, or this protocol can be performed directly after the slide preparation protocol. If performed after RNA FISH, the authors recommend labeling the Cot-1 RNA FISH probe with Cy5-dUTP and using the FITClabeled X-chromosome paint. First, the chromosome paint probe is prepared. Then, the slides are treated to denature the DNA. The probe hybridization and washing steps are identical to those in the RNA FISH protocol.

Additional Materials (also see Basic Protocol 2)

Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

Unlabeled Cot-1 DNA Hybridization buffer (Cambio) Concentrated whole X-chromosome paint (Cambio) 0.2% (v/v) Tween in CMF-PBS 2% (w/v) paraformaldehyde (PFA) in CMF-PBS/0.2% (v/v) Tween RNase A (400 μg/ml in CMF-PBS) CMF-PBS 0.2 N HCl/0.2% (v/v) Tween 70% (v/v) formamide/2× SSC (see recipe for 20× SSC) Fixed slides in 70% ethanol (see Basic Protocol 2) Rubber cement

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Prepare X-chromosome paint probe 1. Vacuum dry or precipitate 40 μg unlabeled Cot-1 DNA for each hybridization spot. 2. For each hybridization spot, resuspend 40 μg dehydrated Cot-1 DNA in 8 μl of hybridization buffer. 3. Add 2 μl of concentrated X-chromosome paint to above Cot-1 hybridization buffer. 4. Denature probe 10 min at 80◦ C. 5. Pre-anneal probe 40 min at 42◦ C.

Denature slides 6. If performing DNA FISH directly after Basic Protocol 1, proceed to step 8. If using slides after RNA FISH (see Basic Protocol 2), remove coverslip and place in a Coplin jar with 0.2% Tween/CMF-PBS. 7. Fix RNA FISH signal by transferring to 2% PFA/CMF-PBS/0.2% Tween. Incubate 10 min at room temperature. 8. Transfer slides to ice -cold 80% ethanol and incubate 5 min. 9. Transfer slides to ice-cold 90% ethanol and incubate 5 min. 10. Transfer slides to ice-cold 100% ethanol and incubate 5 min. 11. Air dry slides. 12. Pipet 50 μl of RNase A (400 μg/ml) onto each hybridization spot. Gently cover the spot with ParaÞlm and incubate in a humidiÞed chamber for 40 min at 37◦ C. 13. Transfer slides to CMF-PBS. 14. Transfer slides to 0.2 N HCl/0.2% Tween. Incubate 10 min at room temperature. 15. Neutralize slides with 0.2%Tween/CMF-PBS for 5 min at room temperature. 16. Denature slides in 70% formamide/2× SSC for 10 min at 80◦ C. 17. Repeat the ethanol dehydration by transferring slides to ice-cold 80% ethanol and incubating for 5 min. 18. Transfer slides to ice-cold 90% ethanol and incubate 5 min. 19. Transfer slides to ice-cold 100% ethanol and incubate 5 min. 20. Air dry slides. 21. Place slides on pre-warmed (37◦ C) humidiÞed chamber. Add 10 μl of pre-annealed probe to the spot of cells on the slides. Carefully cover the probe with a coverslip, avoiding bubbles. Seal the coverslip with rubber cement. 22. Incubate overnight at 37◦ C.

Wash slides 23. Pre-warm three Coplin jars with 2× SSC/50% formamide and three Coplin jars with 2× SSC 20 min at 45◦ C. 24. Remove coverslips. Wash slides three times (5 min each time) in 2× SSC/50% formamide at 45◦ C with light agitation. 25. Wash slides three times (5 min each time) in 2× SSC at 45◦ C with light agitation. 26. Apply 10 μl Vectashield with DAPI mounting medium to each cell spot. Seal coverslips with clear nail polish. Current Protocols in Stem Cell Biology

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27. Visualize slides with an upright epißuorescence microscope with a 60× oil immersion lens or with a confocal microscope equipped with Þlters that are compatible for imaging with DAPI (400 nm), Cy3 (570 nm), Cy5 (660 nm), and FITC (505 nm). Store the slides at 4◦ C in the dark. Signals are stable for a minimum of 6 months. Refer to Silva et al. (2008) for examples of staining patterns.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

CSK-T buffer 100 mM NaCl 200 mM sucrose 10 mM PIPES 3 mM MgCl2 Adjust to pH 6.8 with NaOH Filter sterilze with a 0.2-μm Þlter or autoclave Add Triton X-100 to a Þnal concentration of 0.5% (v/v) Store up to 1 year at 4◦ C Hybridization buffer 50% (w/v) formamide 2× SSC (see recipe for 20× SSC) 2 mg/ml BSA 10% (w/v) dextran sulfate (500 kDa) Store up to 1 year at −20◦ C Mouse embryonic Þbroblast (MEF) medium DMEM-f12 (Gibco, cat. no. 11330) containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock; e.g., Invitrogen) Store up to 4 weeks at 4◦ C SSC, 20× 0.3 M NaCl 0.3 M sodium citrate, pH 7.0 Autoclave Store up to 1 year at room temperature 2× working solution: Dilute 1:10 in DEPC-H2 O. COMMENTARY Background Information

Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

hESC and hPSC can be maintained in culture in a self-renewing state and differentiate to all three embryonic germ layers (Thomson et al., 1998; Takahashi et al., 2007; Yu et al., 2007). While there is great therapeutic potential for these cells, many studies report genetic and epigenetic instabilities during culture (Baker et al., 2007). hPSC laboratories

commonly practice routine karyotypic analysis to monitor genetic instability; however, epigenetic stability is often overlooked. Assaying for XCI in undifferentiated and differentiated cells serves as a useful tool to monitor the epigenetic status of hPSCs. XCI is the epigenetic silencing of an entire X chromosome in female mammals to equalize the gene dosage between XX females

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and XY males (reviewed in Payer and Lee, 2008). XCI is intimately linked to embryonic development, is regulated by the pluripotency factors Oct4, Sox2, and Nanog (Navarro et al., 2008; Donohoe et al., 2009), and is required for female mammalian development. In the mouse, female pre-implantation epiblast cells in the embryo and ES cells in culture have two active X chromosomes and undergo random XCI during differentiation (where either X chromosome is inactivated). However, mouse epiblast stem cells, which are derived from post-implantation epithelialized epiblast, have already undergone XCI, containing one inactive X chromosome expressing XIST (Guo et al., 2009). While the exact timing of XCI within the human epiblast is not known, human embryos at some point have two active X chromosomes and also undergo random XCI during early development. In normal development, once a cell undergoes XCI, the daughter cell inherits and maintains the inactive X chromosome. hESCs in principle have the capacity to recapitulate XCI when induced to differentiate (XIST negative when undifferentiated and XIST positive when differentiated), which can be seen in certain early passage lines (Hall et al., 2008; Silva et al., 2008). Thus, hESCs that undergo XCI upon differentiation could reßect an earlier, less differentiated cell type when compared to hESC, which always express XIST. Due to the epigenetic instability of hESC, several studies report widely disparate XCI statuses, unlike the mouse ESC counterpart (Hoffman et al., 2005; Hall et al., 2008; Shen et al., 2008; Silva et al., 2008). While variation between cell lines could be reßective of the timing of XCI in the human embryo, the majority of disparate XCI states are due to epigenetic instability. Studies report different XCI states even within the same hESC line, in the same laboratory and occasionally within the same culture, demonstrating the epigenetic instability within an hESC line. hESCs fall into three classes of XCI states: upregulating XIST upon differentiation, always expressing XIST in the undifferentiated and differentiated states, and never expressing XIST in the undifferentiated and differentiated states. While certain early passage hESC can recapitulate XCI when induced to differentiate, this status is often the most difÞcult to maintain in culture. These cultures tend to drift into cells that have already undergone XCI in the undifferentiated state. Interestingly, undifferentiated hESC that have undergone XCI can also lose XIST RNA expression by DNA methy-

lation of the XIST promoter while still maintaining an inactive X chromosome (Shen et al., 2008). While these cells do not express XIST, the cells maintain at least a partially inactive X chromosome. The inactive X chromosome is observable by an absence of a Cot-1 signal (Cot-1 hole) co-localizing with one of the two X chromosomes. This last status, where lines lose the ability to up regulate XIST expression upon differentiation, has been found to be the most common, irreversible and clinically problematic. This XCI status is never observed in healthy female adult tissue but is shared by embryonal carcinoma cells and certain malignancies such as breast and ovarian cancer. Several breast and ovarian cancer cells and cell lines lack XIST expression or a fully silenced X chromosome (Pageau et al., 2007). While it is unclear if improper XCI plays a causative role in cancer, misregulated XCI is not observed in healthy adult tissue. (Andrews et al., 2005; Sirchia et al., 2005). Thus, assaying for XCI is an additional tool to characterize and monitor hPSC lines and may have implications for clinical utility and safety. Other epigenetic modiÞcations, such as DNA methylation of imprinted genes, may be useful to monitor epigenetic stability in both male and female ES cells. In general, RT-PCR analysis for XIST will give similar results to FISH analysis, such that cultures that are XIST positive by RT-PCR are also positive by FISH. However, only FISH assays XIST expression on an individual cell level, which can potentially deÞne a mixed population of cells. Also, low levels of XIST expression detected by RT-PCR could reßect a culture in which a small percentage of cells have undergone XCI (visualized by FISH as XIST clouds coating the X chromosome) or a culture with cells before initiating XCI in which all of the cells are expressing low levels of XIST (visualized by FISH as small XIST pinpoints that do not coat an entire X chromosome). The RNA/DNA FISH protocol also assays for X chromosome number (only cells with two X chromosomes in a diploid background will undergo XCI) and assays for the state of silencing by combining Cot-1 FISH with X-chromosome DNA FISH. To date, the status of XCI in human iPSCs has not been reported. However, mouse iPSCs behave like mouse ESCs with two active X chromosomes in the undifferentiated state (XIST negative) and an inactive X chromosome in the differentiated state (XIST positive; Maherali et al., 2007). Thus, the XCI status in human iPSCs may reßect the completeness

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of epigenetic reprogramming. Because human iPSCs share culturing techniques with hESC, they may also share the feature of epigenetic instability. Markers of XCI can be used as one effective screen to monitor the epigenetic status of hPSCs, providing important clinical information on the safety of hPSC therapies.

Critical Parameters and Troubleshooting Probe preparation The DNA used for probe preparation may be from circular plasmids puriÞed from mini or maxi preps or linearized plasmids. Probe quality is highly affected by hybridization buffer quality and reagents must be prepared in an RNase-free manner. Slide preparation Gentle cell trypsinization is essential for successful slide preparation. Excessive background and abnormal Cot-1 staining is often a reßection of harsh treatment during harvesting of cells, which can also be monitored by trypan blue exclusion staining to give an idea of the viability of the culture after harvesting. Accutase (Invitrogen) can also be used to disrupt colonies. Solutions must be RNase-free, and 4% paraformaldehyde in CMF-PBS is stable for only 3 weeks when stored at 4◦ C. RNA signals are only stable on slides for ∼3 weeks when stored at 4◦ C in 70% ethanol. After ethanol dehydration, slides should be transferred to −80◦ C for long-term storage. RNA FISH Hybridization must take place in a humidiÞed chamber to avoid drying of the probe. If intracellular background is too high, increasing the hybridization temperature to 45◦ C can produce a more speciÞc signal. If the signal is weak, slides can be hybridized and washed at 37◦ C. Before moving to sequential RNA and DNA FISH, the RNA FISH signal should be checked and photographed. RNA signals will weaken after DNA FISH is performed; however, the RNA signal is usually still visible. If encountering trouble with RNA signal preservation, the RNA signal can be photographed Þrst with the XY coordinates of the slide recorded. Then DNA FISH can be performed and photographed using the same XY coordinates to overlap the signals. Characterization of X-Chromosome Inactivation Status in Human Pluripotent Stem Cells

DNA FISH Hybridization must take place in a humidiÞed chamber and the coverslips should be sealed with rubber cement. In cases of high

background or weak signal, denaturation conditions or Cot-1 blocking concentration can be adjusted.

Anticipated Results A good in situ hybridization experiment will clearly identify the location of a message within the cells that is speciÞc to inside the nuclei. A positive XIST signal appears as a single cloud that co-localizes with one of the two X chromosomes visualized by X-paint DNA FISH. Cot-1 FISH is a nuclear-wide stain that is absent from the nucleolus but present in all cells. Examples of these stainings can be found in Silva et al. (2008).

Time Considerations Starting from probe labeling, cell harvest, RNA, and sequential DNA FISH, this procedure takes 3 days, with two overnight hybridization steps. As is written in each protocol, probes can be prepared Þrst and stored for 1 year at –20◦ C. After cells are harvested and Þxed, the slides can be stored for 3 weeks at 4◦ C in 70% ethanol or for long-term storage, dehydrated at –80◦ C.

Acknowledgements Work on federally approved hESC lines was funded by a National Institutes of Health grant no. R01-GM58839-S1; work on nonfederally approved hESC lines was funded by a Harvard Stem Cell Institute grant and also by the Howard Hughes Medical Institute.

Literature Cited Andrews, P.W., Matin, M.M., Bahrami, A.R., Damjanov, I., Gokhale, P., and Draper, J.S. 2005. Embryonic stem (ES) cells and embryonal carcinoma (EC) cells: Opposite sides of the same coin. Biochem. Soc. Trans. 33:1526-1530. Baker, D.E., Harrison, N.J., Maltby, E., Smith, K., Moore, H.D., Shaw, P.J., Heath, P.R., Holden, H., and Andrews, P.W. 2007. Adaptation to culture of human embryonic stem cells and oncogenesis in vivo. Nat. Biotechnol. 25:207-215. Donohoe, M.E., Silva, S.S., Pinter, S.F., Xu, N., and Lee, J.T. 2009. The pluripotency factor Oct4 interacts with Ctcf and also controls X-chromosome pairing and counting. Nature 460:128-132. Guo, G., Yang, J., Nichols, J., Hall, J.S., Eyres, I., MansÞeld, W., and Smith, A. 2009. Klf4 reverts developmentally programmed restriction of ground state pluripotency. Development 136:1063-1069. Hall, L.L., Byron, M., Butler, J., Becker, K.A., Nelson, A., Amit, M., Itskovitz-Eldor, J., Stein, J., Stein, G., Ware, C., and Lawrence, J.B. 2008. X-inactivation reveals epigenetic anomalies in

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most HESC but identiÞes sublines that initiate as expected. J. Cell Physiol. 216:445-452.

and prone to epigenetic alterations. Proc. Natl. Acad. Sci. U.S.A. 105:4709-4714.

Hoffman, L.M., Hall, L., Batten, J.L., Young, H., Pardasani, D., Baetge, E.E., Lawrence, J., and Carpenter, M.K. 2005. X-inactivation status varies in human embryonic stem cell lines. Stem Cells 23:1468-1478.

Silva, S.S., Rowntree, R.K., Mekhoubad, S., and Lee, J.T. 2008. X-chromosome inactivation and epigenetic ßuidity in human embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 105:48204825.

Maherali, N., Sridharan, R., Xie, W., Utikal, J., Eminli, S., Arnold, K., Stadtfeld, M., Yachechko, R., Tchieu, J., Jaenisch, R., Plath, K., and Hochedlinger, K. 2007. Directly reprogrammed Þbroblasts show global epigenetic remodeling and widespread tissue contribution. Cell Stem Cell. 1:55-70.

Sirchia, S.M., Ramoscelli, L., Grati, F.R., Barbera, F., Coradini, D., Rossella, F., Porta, G., Lesma, E., Ruggeri, A., Radice, P., Simoni, G., and Miozzo, M. 2005. Loss of the inactive X chromosome and replication of the active X in BRCA1-defective and wild-type breast cancer cells. Cancer Res. 65:2139-2146.

Navarro, P., Chambers, I., Karwacki-Neisius, V., Chureau, C., Morey, C., Rougeulle, C., and Avner, P. 2008. Molecular coupling of XIST regulation and pluripotency. Science 321:16931695.

Takahashi, K., Tanabe, K., Ohnuki, M., Narita, M., Ichisaka, T., Tomoda, K., and Yamanaka, S. 2007. Induction of pluripotent stem cells from adult human Þbroblasts by deÞned factors. Cell 131:861-872.

Pageau, G., Hall, L., Ganesan, S., Livingston, D., and Lawrence, J. 2007. The disappearing Barr body in breast and ovarian cancers. Nat. Rev. Cancer 7:628-633.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

Payer, B. and Lee, J.T. 2008. X chromosome dosage compensation: How mammals keep the balance. Annu. Rev. Genet. 42:733-772. Shen, Y., Matsuno, Y., Fouse, S.D., Rao, N., Root, S., Xu, R., Pellegrini, M., Riggs, A.D., and Fan, G. 2008. X-inactivation in female human embryonic stem cells is in a nonrandom pattern

Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S., Nie, J., Jonsdottir, G.A., Ruotti, V., Stewart, R., Slukvin, II, and Thomson, J.A. 2007. Induced pluripotent stem cell lines derived from human somatic cells. Science 318:1917-1920.

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Preparation of Defined Human Embryonic Stem Cell Populations for Transcriptional Profiling

UNIT 1B.7

Qi Zhou,1 Hun Chy,1 and Andrew L. Laslett1,2 1

Commonwealth Scientific and Industrial Research Organization (CSIRO), Molecular and Health Technologies, Clayton, Australia 2 Department of Anatomy and Developmental Biology, Monash University, Clayton, Australia

ABSTRACT This unit describes a useful approach to preparing highly reproducible samples of human embryonic stem cell (hESC) total RNA suitable for transcriptional profiling from heterogeneous mixtures of cells containing undifferentiated hESC and differentiated cell types. In this unit, fluorescence-activated cell sorting (FACS) is used to sub-fractionate hESC populations on the basis of their levels of co-expression of two previously published hESC surface markers, CD9(TG30) and GCTM-2. This sub-fractionation allows for the separation of undifferentiated hESC (CD9hi, GCTM-2hi) from the early stages in hESC differentiation (CD9neg or low, GCTM-2neg or low). Curr. Protoc. Stem Cell C 2010 by John Wiley & Sons, Inc. Biol. 14:1B.7.1-1B.7.15.  Keywords: human embryonic stem cells r cell surface markers r fluorescence-activated cell sorting

INTRODUCTION This unit describes an easily reproducible protocol for the preparation of high-quality RNA samples useful for large-scale gene expression analyses. The unit begins with a description of a protocol for the routine maintenance of hESC (Basic Protocol 1), although the subsequent protocols are applicable to hESC cultured in a variety of different conditions. The following sections describe detailed protocols for immunofluorescent labeling of hESC (Basic Protocol 2), FACS separation (Basic Protocol 3, and RNA isolation (Basic Protocol 4).

ENZYMATIC PASSAGING OF CULTURES OF hESCs This method can be used to passage iPS or hESC grown in different media and on various feeders and matrices.

BASIC PROTOCOL 1

Materials hESCs in culture Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) Collagenase I solution (4 mg/ml in DMEM/F-12) Knock-out serum replacement (KSR) medium (see recipe) Mouse embryonic fibroblast (MEF) feeder cells (Pera et al., 2003; UNIT 1C.3) plated on gelatinized flasks at 1.2 × 104 cells/cm2 Inverted microscope Dissecting microscope 37◦ C incubator 10-ml pipet

Current Protocols in Stem Cell Biology 1B.7.1-1B.7.15 Published online July 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01b07s14 C 2010 John Wiley & Sons, Inc. Copyright 

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feeder cells

differentiated cells

undifferentiated cells

Figure 1B.7.1 Bright-field image of a day 7 hESC colony cultured on mouse embryonic fibroblasts that is ready to be passaged by treatment with collagenase (50×). Note the compact appearance of undifferentiated hESC in the center of the colony, surrounded by a ring of differentiated cells.

Cell scraper, optional 15-ml tubes Centrifuge Prepare the cultures 1. On the day of planned passage, check the hESC culture under a microscope and evaluate the colonies for suitability of passaging (see Fig. 1B.7.1). 2. Aspirate the medium and wash the cells twice with an appropriate volume of prewarmed (37◦ C) CMF-PBS (e.g., 10 ml for a 75-cm2 flask).

Digest the culture 3. Aspirate the CMF-PBS and add collagenase I solution (e.g., 1, 3, and 7 ml for 25cm2 , 75-cm2 , and 175-cm2 flask, respectively). Incubate at 37◦ C until the colonies begin to curl around the edges, as observed using an inverted microscope. As a guide, it can take ∼5 min to observe this for feeder-free cultures and 6 min for those on feeders.

4. Aspirate the collagenase I solution.

Collect the cells 5. Pipet fresh KSR medium into the flask (3 or 7 ml for 25-cm2 and 75-cm2 , respectively), and wash down the flask wall to dislodge hESC colonies. If necessary, use the tip of the 10-ml pipet or a cell scraper to scrape off colonies.

Preparation of Defined hESC Populations for Transcriptional Profiling

6. Collect the cell suspension into the pipet and wash down the flask wall again, repeat as necessary to collect all of the cells, and break up the colonies into small clumps of ∼10 to 100 cells. 7. Transfer the cell suspension into a 15-ml tube. Add an additional 3 or 7 ml (25-cm2 and 75-cm2 , respectively) fresh KSR medium into the flask and wash down the flask wall 3 to 4 times to dislodge the remaining hESC colonies

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8. Transfer this cell suspension to the same 15-ml tube and gently resuspend the cell clumps, before letting the cells settle for exactly 5 min. If the cultures were highly differentiated, reduce the settling time to 1 to 2 min. This allows clumps of hESC to settle by gravity and separate from the majority of the MEF feeder cells that stay in suspension. In our experience, the MEF feeder cells become a single-cell suspension as they are lifted from the flask after collagenase treatment. This strategy, in our hands, drastically reduces contamination by MEF feeder cells.

9. Aspirate the top half volume of the cell suspension to remove any differentiated hESC and fibroblast feeder cells, which are lighter and will remain in the top half of the suspension during this short settling period.

Plate the cells 10. Centrifuge the remaining cell suspension 2 min at 170 × g, 4◦ C, and determine the final split-ratio from the size of the cell pellet. Initial split ratios should be conservative (e.g., 1:1 or 1:2) and operators need to empirically determine the appropriate split ratio based on the confluency of each flask and the desired passaging cycle.

11. Aspirate the supernatant and resuspend the cell pellet in appropriate volume of medium according to the split-ratio. (e.g., if applying a 1:8 split ratio, add 8 ml of KSR medium, resuspend the cells, and use 1 ml of cell suspension for each flask to be plated). For example, one confluent 25-cm2 flask split into three 75-cm2 flasks is approximately a 1:9 split.

12. Plate the appropriate number of cells onto a gelatinized flask containing mitotically inactivated MEFs seeded previously at a density of 1.2 × 104 cells/cm2 , agitate them controllably in a backwards/forwards and side-to-side motion to distribute the cells evenly, and transfer the flasks into the incubator. 13. Change the KSR medium daily until the colonies are ready to be passaged again. Over the weekends and holidays, it is sufficient to change the medium only once every 2 days, e.g., either Saturday or Sunday).

STARTING hESC BULK CULTURE FROM MAINTENANCE CULTURE The following method outlines the procedure for initiating enzymatically passaged “bulk” cultures from mechanically passaged stem cell cultures (Pera et al., 2003) The protocol is designed for initiating a bulk culture in 25-cm2 format starting from two organ culture dishes. In order to reduce the probability of acquired karyotypic abnormalities in enzymatically passaged cultures, we initiate fresh bulk cultures from our maintenance cultures every 8 to 12 weeks. Large batches of bulk cultures may be cryopreserved at low passage number.

ALTERNATE PROTOCOL 1

Materials Mechanically passaged hESCs in organ culture dishes PBS with Ca++ and Mg++ (PBS+ ) Dispase solution (10 mg/ml in hESC medium; see recipe), ice cold Gelatinized 25-cm2 flasks seeded with 1.2 × 104 cells/cm2 inactivated MEFs (UNIT 1C.3) Knock-out serum replacement (KSR) medium (see recipe) Dissecting microscope with warm stage 4-well culture dish (Nunc, cat. no. 176740) Pulled glass capillary cutter or a 26-G needle attached to a 3-ml syringe or a sterilized ultra sharp splitting blade Current Protocols in Stem Cell Biology

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Micropipet tips 20-μl or 200-μl micropipet tip 1.5-ml microcentrifuge tubes Evaluate the colonies 1. On the day of passaging, check hESC colonies to be passaged under a microscope to ensure suitability for passage. The authors use colonies from two organ culture dishes for each 25-cm2 flask.

2. Prepare a 4-well dish with 1 ml PBS+ in each well and warm it at 37◦ C.

Dissect undifferentiated areas and digest 3. Manually cut around whole colonies and cut out differentiated parts of the colonies including the central button using either a sterile prepulled glass capillary pipet attached to a capillary holder, a 26-G needle attached to a 3-ml syringe, or a cleaned and sterilized ultra sharp splitting blade (see Fig. 1B.7.2). 4. Remove the central buttons and differentiated pieces with a micropipet tip. 5. When all of the colonies on the organ culture dish have been cut and differentiated parts removed, aspirate the medium from the dish, add 500 μl cold dispase solution to each dish, and place the dishes on the warm stage for 2 to 3 min.

Collect undifferentiated fragments 6. When feeder cells have disintegrated, gently nudge off cut pieces and transfer them into the first well of the 4-well PBS+ plate, using either a 20-μl or a 200-μl micropipet tip. 7. Pick up all colonies and serially wash each colony by transferring them from well to well of PBS+ solution until all colonies are in well 4. 8. Pick up all colonies in 100 μl volume of PBS+ and transfer to a sterile 1.5-ml microcentrifuge tube. Gently pipet up and down ∼5 times to break up the colonies, taking care not to break them up too much.

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Figure 1B.7.2 Dark-field image showing a day 7 colony prior to being transferred into bulk passage. The colony has been cut into six pieces. The largest piece has differentiated and the other five pieces are “good” pieces, with minimal visible differentiation, to be transferred into bulk culture. Current Protocols in Stem Cell Biology

Plate the cell fragments 9. Add the entire contents of the microcentrifuge tube onto a gelatinized 25-cm2 flask containing mitotically inactivated MEFs seeded previously at a density of 1.2 × 104 cells/cm2 , gently rock the flask to evenly distribute the hESC colony pieces, and place the flask in incubator. 10. Change the medium daily with KSR medium until the colonies are ready to be passaged again. Once the cells have reached ∼70% confluency after 7 days, passage the cells following the steps in Basic Protocol 1. Over the weekends, it is sufficient to change the medium only once, either Saturday or Sunday.

HARVESTING hESCs FROM BULK CULTURES Enzymatically adapted hESC are harvested using TrypLE Express, a recombinant enzyme similar to porcine trypsin, in order to obtain a high yield of viable cells with an intact surface membrane. Other harvesting methods can be used, but care needs to be taken that enzymatic treatments do not affect the presence of or damage protein epitopes on the surface of hESC.

BASIC PROTOCOL 2

Materials hESCs in bulk cultures (Basic Protocol 1) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) TrypLE Express (Invitrogen, cat. no. 12604) 20% (v/v) fetal bovine serum (FBS; Invitrogen, cat. no. 16000-044) in DMEM/F12 medium (chilled to 4◦ C) 37◦ C incubator 35-μm cell strainer (BD Biosciences, cat. no. 352235) Centrifuge Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3) 1. Aspirate the KSR medium from 75-cm2 flasks (bulk cultures). 2. Wash cells twice, each time with 10 ml CMF-PBS in the flasks, and aspirate the CMF-PBS after each wash. 3. Add TrypLE Express into the flask (e.g., 0.5 or 1 ml for 25-cm2 and 75-cm2 flasks, respectively) and incubate 5 min at 37◦ C. 4. Pipet a few milliliters of 20% FBS/DMEM/F12 medium into the flask to inactivate the TrypLE Express and wash the flask wall to dislodge hESC colonies. Collect the cell suspension into the pipet and wash down the flask wall again, repeat the process a few more times. 5. Pass the cell suspension through a 35-μm cell strainer to generate a single-cell suspension and centrifuge 2 min at 170 × g, 4◦ C. 6. Aspirate the supernatant and resuspend cells in 10 ml 20% FBS/DMEM/F12 medium. Centrifuge 2 min at 170 × g, 4◦ C. 7. Aspirate the supernatant and resuspend cells in 10 ml 20% FBS/DMEM/F12 medium. 8. Take small aliquots of cells for cell number counting using a hemacytometer (UNIT 1C.3). The cells are now ready for immunofluorescent staining. From a 75-cm2 flask, you should expect to recover ∼15 to 20 million cells.

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BASIC PROTOCOL 3

IMMUNOFLUORESCENT STAINING OF SURFACE ANTIGENS ON LIVE hESCs FOR FACS Currently, there are a limited number of cell surface antigens that can be used as reliable markers for undifferentiated hESC that switch off rapidly when hESC differentiate (Laslett et al., 2003). We have previously demonstrated that the use of single surface markers to detect and separate hESC from differentiated cell types is not as efficient as using a combination of cell surface markers (Laslett et al., 2007). Our previous studies utilized antibodies that detect the cell surface antigens GCTM-2 and CD9, both of which down regulate upon hESC differentiation, and demonstrated that co-expression of high levels of immunoreactivity of these markers directly corresponds with enriched populations of hESC (Laslett et al., 2007; Hough et al., 2009; Kolle et al., 2009). The protocol described below for immunofluorescent staining of hESC with antibodies to CD9 and GCTM-2 could also be carried out using other cell surface markers that detect hESC (e.g., SSEA-3 and Tra 1-60). See Table 1B.7.1 for samples for FACS sorting.

Materials Harvested bulk hESCs (Basic Protocol 2) Mouse anti-CD9 (TG30) monoclonal antibody (Millipore, cat. no. MAB4427) Mouse anti-GCTM2 monoclonal antibody (0.4 mg/ml; kind gift from Martin Pera)—a commercially available alternative, which detects the same antigen is the antibody TG343 (Millipore, cat. no. MAB4346) PE rat anti-mouse CD 90.2 (0.2 mg/ml; BD Pharmingen, cat. no. 553014) Mouse IgG2a isotype control (0.5 mg/ml; BD Pharmingen, cat. no. 554121) Mouse IgM isotype control (0.5 mg/ml; BD Pharmingen, cat. no. 553472) 20% (v/v) fetal bovine serum (FBS) in DMEM/F12 medium (chilled to 4◦ C) Alexa Fluor 488 goat anti–mouse IgG2a (2 mg/ml; Invitrogen, cat. no. A21131) Alexa Fluor 647 goat anti–mouse IgM (2 mg/ml; Invitrogen, cat. no. A21238) R-phycoerythrin goat anti–mouse IgG2a conjugate (1.0 mg/ml; Invitrogen, cat. no. P21139) Propidium iodide solution (1.0 mg/ml; Sigma, cat. no. P4864) Spherotech 8 peak Ultra Rainbow beads Centrifuge 35-μm cell strainer (BD Biosciences, cat. no. 352235) FACSVantage DiVa (Becton Dickinson) or equivalent NOTE: Both primary and secondary antibody staining and washing are carried out in 20% FBS medium prechilled to 4◦ C. For 1.5 × 107 cells (recommended cell number for test stainings; fewer cells can be used in control tubes, e.g., 50,000 cells/tube); 10 ml and 2 ml medium are used for primary and secondary antibody staining respectively.

Stain cells for sorting 1. Resuspend the cells in 10 ml primary antibody solution and incubate on ice for 30 min (Table 1B.7.2). hESCs are fragile and must be handled gently. For the single-color controls for CD9(TG30) or GCTM2, the same concentration of each primary antibody should be used as in the double-staining sample. For double staining, the PE rat anti-mouse CD 90.2 can be done together with secondary antibodies staining. Preparation of Defined hESC Populations for Transcriptional Profiling

For mouse immunoglobulin isotype control, the concentration of the two subtype specific immunoglobulins should be equivalent to the two primary antibodies in regard to protein concentration.

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Table 1B.7.1 Samples for FACS Sortinga,b

Sample

Primary antibody

Secondary antibody

PE rat anti-mouse CD 90.2c

Propidium iodide

Double staining for TG30 + GCTM2d sorting

AF 488 goat anti–mouse IgG2a + AF647 goat anti-mouse IgM

+

+

TG30 single-color control

AF 488 goat anti–mouse IgG2a

−

+

AF647 goat anti-mouse IgM

−

+

TG30

GCTM2 GCTM2 single-color control Anti-mouse CD 90.2 control

−

−

+

+

Mouse immunoglobulin isotype control

Mouse IgG2a isotype control + IgM isotype controld

AF 488 goat anti–mouse IgG2a + AF647 goat anti–mouse IgM

−

+

Phycoerythrin control

Mouse IgG2a isotype control

R-phycoerythrin goat anti–mouse IgG2a

−

+

Unstained control

−

−

−

+

a +; added b −; not added c PE rat anti–mouse CD 90.2 was used to specifically recognize mouse cells, thereby removing MEF feeder cells from hESC by FACS (Filipczyk et al., 2007). d Note these samples are stained simultaneously with both antibodies, as indicated.

Table 1B.7.2 Antibody Information for FACS

Antibody

Host

Isotype

Working dilution

CD9(TG30)

Mouse

IgG2a

1:1000

GCTM2

Mouse

IgM

1:50∼1:500a

Rat

/b

1:100

Goat

/

b

1:500

/

b

1:1000

b

1:2000

PE rat anti–mouse CD 90.2 AF 488 goat anti–mouse IgG2a AF647 goat anti–mouse IgM R-phycoerythrin goat anti–mouse IgG2a

Goat Goat

/

a The epitope of GCTM2 is expressed on hESC surface with such extremely high abundance that saturated binding

of monoclonal antibodies is unattainable at feasible concentrations. Therefore, before each new batch of antibody comes into use, a titration experiment against a standard control or the previous batch should be done so that one constant ratio of antibody activity/cell number in different experiments is guaranteed. b Forward slash (/) indicates that these antibodies are polyclonal.

2. Centrifuge the cells 2 min at 170 × g, 4◦ C, aspirate the supernatant, and resuspend the cells in 10 ml 20% FBS/DMEM/F12 medium. 3. Repeat step 2 for one more wash. 4. Resuspend the cells in 2 ml secondary antibody solution and incubate on ice for 30 min (Table 1B.7.2). 5. Centrifuge the cells 2 min at 170 × g, 4◦ C, aspirate the supernatant, and resuspend the cells in 10 ml 20% FBS/DMEM/F12 medium. 6. Repeat step 5 for one more wash. 7. Resuspend the cells in 10 ml 20% FBS/DMEM/F12 medium.

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Strain the cells, and stain dead cells 8. Just before FACS, pass the cell suspension through a 35-μm cell strainer to generate a single-cell suspension. The final filtration of single cells should be done just before FACS, as clumps might form again with the cells sitting on bench.

9. Add propidium iodide to the final concentration of 0.3 μg/ml for nuclear staining of dead cells. The cells are now ready for FACS.

105

Mel1-Sort-Single Cells

100 150 FSC-A

200

250 ( 1,000)

Pl FL3-A 103 104 50

100 150 FSC-H

Mel1-Sort-P1

200

1,097

50 SSC-A ( 1,000) 100 150 200 250

50

250 ( 1,000)

50

100 150 FSC-A

200

250 ( 1,000)

105

Mel1-Sort-P1 P7

3

0

103

104

FL2-A

105

GCTM2-A647 FL8-A 286 104 102 0 102 103

P6

50 10 1,890

Mel1-Sort Single Cells

50

Mel1-Sort-Single Cells

0

FSC-A ( 1,000) 100 150 200 250

SSC-A ( 1,000) 100 150 200 250

A typical FACS plot demonstrating staining for CD9 and GCTM2 is shown in Figure 1B.7.3.

163

P5

P4

0

102

104 103 TG30-A488 FL1-A

Experiment Name:

DGZ071114

Tube: Sort

Specimen Name:

Mel1

Tube Name:

Sort

Record Date:

Nov 14, 2007 12:00:0...

# Events % Parent % Total Population 100.0 100,000 All Events 95.8 95.8 95,833 Single Cells 85.9 89.6 85,868 P1 85.2 99.2 85,172 P2 2.4 2.8 2,364 P4 25.4 29.8 25,398 P5 34.2 40.1 34,151 P6 8.7 10.3 8,747 P7

Population

105

Figure 1B.7.3 Typical sorting strategy for hESCs. Sorted cells were initially gated using forward and side scatter, followed by the removal of clumps and doublets by gating on single cells (FSC-A vs. FSC-H), nonviable cells were excluded based on propidium iodide (PI) fluorescence and, penultimately, MEF feeder cells were removed using negative selection for Thy1.2-PE. hESC were then separated by FACS on the basis of cell surface intensity of GCTM-2 and TG30 (CD9) into four populations: P4 (GCTM-2neg CD9neg ), P5 (GCTM-2low CD9low ), P6 (GCTM-2mid CD9mid ), and P7 (GCTM-2high CD9high ). The P4 gate is set relative to isotype controls and the P5-P7 gates are reproducibly set based on initial experiments, using the strategy outlined in Figure 1B.7.4.

1B.7.8 Supplement 14

Current Protocols in Stem Cell Biology

FACSDiva Version 6.1.1

100

SSC-A

150

200

( 1000) 250

HES-Rainbow

50

P1

50

100

150

200

FSC-A

250 ( 1000)

Count 0 10 20 30 40 50 60 70 80 90

HES-Rainbow

P2

102

103

FL1-A

104

105

0

Count 10 20 30 40 50 60 70

HES-Rainbow

P3

102

103

FL2-A

104

105

Count 0 5 10 15 20 25 30 35 40 45

HES-Rainbow

P4

102

103

FL8-A

104

105

Figure 1B.7.4 The template used to set up the FACS DIVA prior to each FACS experiment. Spherotech beads were initially gated using forward and side scatter to exclude debris. The FACS DIVA was then calibrated so that beads of known fluorescence were always gated into the same preset gates (P2 for FL1, P3 for FL2, and P4 for FL8). This ensures that the FACS DIVA is identically calibrated for each separate experiment. Embryonic and Extraembryonic Stem Cells

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Standardize fluorescence detection 10. Prior to running each experiment, use rainbow calibration particles to set the sensitivity of the fluorescence detectors of the flow cytometer. This is done to ensure that the detected fluorescence emission (and the resulting distribution of sorted cells) is comparable between experiments conducted on different days, and also that the results are not influenced by fluctuations in the daily calibration settings of the cell sorter. In these experiments, samples were sorted on a FACSVantage DiVa (Becton Dickinson).

11. Before the control and sample tubes are run, run Spherotech 8 peak Ultra Rainbow beads in a preset histogram template that is created at the beginning of the overall series of experiments (Fig. 1B.7.4). The template includes a histogram for each fluorescence parameter to be measured; FL1 (Alexa Fluor 488), FL2 (PE), and FL8 (Alexa Fluor 647), and each histogram contains a preset gate that coincides with the desired position of the second brightest peak of the calibration beads.

12. During bead acquisition, adjust the gain for each fluorescence parameter to position the second highest peak of the rainbow beads in the center of the preset gate. 13. Use these settings to run the control and sample tubes in the experimental template. 14. Sort the cells into 1.5-ml microcentrifuge tubes or 12-well plates using the FACS Vantage DIVA. Initially gate-sorted cells were using forward and side scatter, followed by the removal of clumps and doublets by gating on single cells (forward scatter [FSC]-A vs. FSC-H), and removal of MEF feeder cells using negative selection for Thy1.2-PE (Fig. 1B.7.3). ALTERNATE PROTOCOL 2

COMBINED DETECTION OF OCT-4 INTRACELLULAR EXPRESSION AND OTHER CELL SURFACE MARKERS ON FIXED hESC BY FLOW CYTOMETRY The nuclear transcription factor OCT-4 is one of the most widely utilized markers of the undifferentiated ESC state (Boyer et al., 2005). OCT-4 forms an elaborate autoregulatory network with the transcription factors SOX2 and NANOG (Chambers et al., 2003; Mitsui et al., 2003). In order to directly assess the correlation between OCT-4 and other related proteins, it is essential to simultaneously detect the expression of OCT-4 and related proteins on individual cells by flow cytometry. As OCT-4 is an intracellular (nuclear) protein, cell permeabilization is required for its detection by flow cytometry, making this protocol incompatible with the recovery of live cells. However, the following method allows simultaneous staining for OCT-4 and surface markers.

Preparation of Defined hESC Populations for Transcriptional Profiling

Fixation/permeabilization treatment of cells exposes all cytosolic proteins to both primary and secondary antibodies, resulting in much higher background due to nonspecific binding of antibodies. This method performs the staining of cell surface markers first, prior to fixation/permeabilization and final staining of intracellular markers, avoiding some of the nonspecific binding of the surface antibodies to the cytosolic components and possible disruption of surface antigens by fixation/permeabilization treatment. According to our experience, however, background for such intracellular staining is still quite high. Therefore, it is important to use a blocking agent (goat serum in this case) for all intracellular staining and, as with all flow cytometric protocols, care must be taken that compensation settings are set appropriately.

1B.7.10 Supplement 14

Current Protocols in Stem Cell Biology

C High 90%

100

103 hi

TG343 AF647

102

101

102

54% E

39%

101 medium 100

103

70%

Medium

7% negative/low 100

101 102 TG30 (CD9) AF488

100

103

101

102

103

G Negative/low 22%

100

101

102

103

OCT4 R-PE

Figure 1B.7.5 Data from a representative flow cytometry analysis of hESC using TG343, CD9, and OCT4. hESC were initially gated using forward and side scatter, followed by the removal of clumps and doublets by gating on single cells (FSC-A vs. FSC-H), nonviable cells were excluded based on propidium iodide (PI) fluorescence and, penultimately, MEF feeder cells were removed using negative selection for Thy1.2-PE (not shown). hESC were then analyzed using the FACS DIVA for immunoreactivity to TG343, CD9, and OCT4. TG343hi TG30hi cells were 90% positive for OCT-4, TG343medium TG30medium cells were 70% positive for OCT-4, and TG343negative/low TG30negative/low cells were 22% positive for OCT4.

Note also that due to the permeabilization step, all cells will take up PI, which in this case cannot be used to give an indication of cell viability. Figure 1B.7.5 shows data from a representative flow cytometric analysis of hESC using TG343, CD9, and OCT4.

Materials Harvested hESCs (Basic Protocol 2) 2% (w/v) paraformaldehyde (PFA) solution in CMF-PBS, pH 7.4 10% (v/v) goat serum in CMF-PBS Phosphate-buffered serum, calcium- and magnesium-free (CMF-PBS) 0.1% (v/v) Triton X-100 solution in CMF-PBS Mouse anti-Oct4 monoclonal antibody(1 mg/ml; Chemicon, cat. no. MAB4401) R-phycoerythrin goat anti-mouse IgG1 conjugate (1 mg/ml; Invitrogen, cat. no. P21129) Mouse IgG1 isotype control (0.5 mg/ml; BD Pharmingen, cat. no. 553447) Centrifuge 35-μm cell strainer (BD Biosciences, cat. no. 352235)

Embryonic and Extraembryonic Stem Cells

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Fix the cells 1. Using 1 × 106 hESCs, perform surface antigen staining (see Basic Protocol 3, steps 1 to 6) in a 1.5-ml microcentrifuge tube. 2. Resuspend cells in 1 ml of 2% PFA, leave at room temperature for 30 min with occasional mixing. 3. Wash the sample in 1 ml 10% goat serum, pellet cells by centrifuging 5 min at 170 × g, room temperature.

Permeabilize the cells 4. Resuspend the cells in 1 ml 0.1% Triton X-100, leave at room temperature for 5 min. 5. Wash twice, each time in 1 ml 10% goat serum.

Block the cells 6. Resuspend the cells in 1 ml 10% goat serum, leave at room temperature for 30 min for blocking. The cells are now ready for intracellular staining. Use 10% goat serum in all subsequent antibody incubation steps.

Stain intracellular antigens 7. Resuspend cells in 1 ml primary antibody solution at the appropriate dilution (Table 1B.7.3) and incubate for 30 min on ice. 8. Centrifuge the cells 2 min at 170 × g, room temperature, aspirate the supernatant, and very gently resuspend the cells in 1 ml 10% goat serum. 9. Repeat step 8 for one more wash. 10. Resuspend cells in 1 ml secondary antibody solution at the appropriate dilution (Table 1B.7.3) and incubate for 30 min on ice. 11. Centrifuge the cells 2 min at 170 × g, room temperature, aspirate the supernatant, and very gently resuspend the cells in 1 ml 10% goat serum. 12. Repeat step 11 for one more wash.

Visualize the staining 13. Resuspend the cells in 1 ml CMF-PBS and pass the cell suspension through a 35-μm cell strainer. The cells are now ready for flow cytometry. Figure 1B.7.5 shows data from a representative flow cytometric analysis of hESC with TG343, CD9, and OCT4 using a FACSVantage DiVa. Table 1B.7.3 Antibody Information for Combined Intracellular and Cell Surface Immunodetection

Antibody Mouse anti-Oct-4 monoclonal antibody R-phycoerythrin goat anti–mouse IgG1

Host

Subtype

Working dilution

Mouse

IgG1

1:200

Goat

a

1:2000

/

a Forward slash (/) indicates that these antibodies are polyclonal.

Preparation of Defined hESC Populations for Transcriptional Profiling

1B.7.12 Supplement 14

Current Protocols in Stem Cell Biology

RNA ISOLATION The RNA isolation and purification technique is largely determined by individual laboratory preferences and the availability of a quick and easy procedure. Cell number is also a major factor, which determines the choice of RNA extraction procedure and the kit to use. Numerous RNA extraction procedures and kits have been described by both laboratory groups and manufacturers. Here, we briefly describe our preferred protocol to generate RNA and cDNA of the desired quality for microarray analysis. The high quality cDNA can also be used for polymerase chain reaction (PCR) amplification and quantitative real-time PCR.

BASIC PROTOCOL 4

Materials hESC bulk cultures (Basic Protocol 2) RNeasy Mini kit (Qiagen, cat. no. 74104) RNase-free H2 O Nanodrop ND 1000 spectrophotometer (Thermo Fisher Scientific) RNase-free tubes (PCR tubes) NOTE: Keep all reagents, working solutions, and master mixes on ice during these procedures.

Purify RNA 1. Harvest hES cells (as described in Basic Protocol 2) and purify total RNA using the Qiagen RNeasy mini kit per manufacturer’s instructions (Protocol: Purification of Total RNA from Animal Cells Using Spin Technology). 2. Use the Nanodrop ND 1000 spectrophotometer to determine the quantity and purity of the extracted RNA. RNA quality is acceptable if the ratio of absorbance at 260 nm/280 nm is between 1.8 and 2.0.

3. Calculate the concentration of RNA and aliquot 2 μg into a RNase-free tube. 4. Resuspend the 2 μg of RNA in 100 μl of RNase-free water. Total RNA generated should be stored at −80◦ C until required for use in microarray experiments. For their studies, the authors then sent the total RNA to a commercial array facility for cDNA synthesis.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

hESC medium DMEM with high glucose (Invitrogen, cat. no. 11960-044) containing: Fetal bovine serum (FBS; 20% v/v final concentration) Nonessential amino acids (Invitrogen, cat. no. 11140-050; 0.1 mM final concentration) Insulin-transferrin-selenium (Invitrogen, cat. no. 41400-045; 1× final concentration) 2-mercaptoethanol (Invitrogen, cat. no. 21985-023; 90 μM final concentration) continued

Embryonic and Extraembryonic Stem Cells

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Supplement 14

50 U/ml penicillin/50 μg/ml streptomycin (from a stock solution of 5000 U/ml penicillin G and 5000 μg/ml streptomycin sulfate; Invitrogen, cat. no. 15070063) Filter sterilize using a 0.22-μm pore size filter (Millipore) Store up to 1 month at 4◦ C Knock-out serum replacement (KSR) medium DMEM/F12 (Invitrogen, cat. no. 11320-033) containing: Knockout serum replacement (Invitrogen, cat. no. 10828-028; 20% v/v final concentration) Nonessential amino acids (Invitrogen, cat. no. 11140-050; 0.1 mM final concentration) L-glutamine (Invitrogen, cat. no. 25030-081; 1% final concentration) 2-mercaptoethanol (Invitrogen, cat. no. 21985-023; 90 μM final concentration) Store up to 1 month at 4◦ C For culturing purpose, human basic fibroblast growth factor (bFGF) needs to be added freshly to 4 ng/ml before use. For washing purpose, do not add bFGF.

COMMENTARY Background Information

Preparation of Defined hESC Populations for Transcriptional Profiling

Human embryonic stem cell (hESC) cultures are composed of a heterogeneous population of cells, some of which have embarked on the pathway to differentiation. Therefore, any study that treats hESC cultures as a homogeneous population of cells is potentially providing only limited insight into the control of stem cell renewal and differentiation. To avoid the problem of dealing with heterogeneous populations of cells, the obvious approach is to use antibodies that detect specific protein markers on the surface of hESC to separate hESC from differentiated cell types. Currently, none of the most commonly used hESC markers are entirely specific for undifferentiated hESC, and immunoreactivity can be observed in embryonic tissues and/or more mature cell types. Therefore, these markers are only useful within the specific context of embryonic stem cell commitment and early differentiation. We have previously demonstrated that co-expression of high levels of monoclonal antibodies to CD9 (TG30) and GCTM-2 correlates with the presence of OCT-4 protein and with large numbers of mRNAs associated with pluripotency (Laslett et al., 2007; Kolle et al., 2009; Hough et al., 2009). The CD9 (TG30) antibody reacts with a cell surface epitope on a 25-kDa protein identified as the tetraspanin protein CD9, while the GCTM-2 antibody recognizes an epitope on the protein core of a high-molecular-weight pericellular matrix keratan sulfate/chondroitin sulfate proteoglycan. Both of these markers are expressed on undifferentiated hESC, down regulate upon

differentiation (Laslett et al., 2007), and in combination can be used to separate hESC from differentiating cells. This can also be achieved by using combinations of other cell surface markers found on hESC (e.g., SSEA3 and Tra 1-60).

Critical Parameters and Troubleshooting The protocols described provide a detailed methodology for the production of good quality RNA from hESC that have been defined using two cell surface markers. When carrying out the procedure it is essential that the gates set on the FACSVantage DiVa, or an alternative FACS sorter, are set identically for each individual experiment. This should be carried out using the Spherotech 8 peak Ultra Rainbow beads run in the preset histogram template that was created at the beginning of the overall series of experiments (see Fig. 1B.7.4). Further troubleshooting tips are outlined in Table 1B.7.4.

Anticipated Results A typical example of the results for FACS separation of hESC is shown in Figure 1B.7.3. A standard experiment using a 75-cm2 flask of 80% confluent hESC would then generate at least 100,000 cells per sorted fraction and subsequently generate at least 100 ng of total RNA/fraction with an absorbance ratio (A260 /A280 ) of ∼2.0, which is the minimum amount of RNA required for use with the Illumina microarray platform. For our studies, we then sent total RNA to a commercial array

1B.7.14 Supplement 14

Current Protocols in Stem Cell Biology

Table 1B.7.4 Troubleshooting Guide to Preparation of hESCs for Transcriptional Profiling

Problem

Possible cause

Solution

Down shift of immunostaining pattern with TG30 and GCTM2 in flow cytometry resulting in very few OR no cells in P7 gate

The culture is differentiated

Repeat the experiment on a new batch of hESCs with microscopically normal look

The titer of the GCTM2 antibody dropped or is over diluted

Repeat the experiment on a new batch of hESCs with microscopically normal look

Low RNA yield

Not enough cells processed for RNA isolation, especially cells from P4 gate whose number can be overestimated

Increase the number of cells used in the experiment

DNA contamination of RNA

No DNase treatment

Perform optional on-column DNase digestion using the RNase-Free DNase Set from the RNeasy Mini Kit

facility, which subjected the RNA to reverse transcription, second-strand cDNA synthesis, and in vitro transcription using the Total Prep RNA Amplification kit (Ambion), followed by hybridization to Illumina Sentrix Human 6 V2 BeadChip arrays (Illumina).

Time Considerations After each enzymatic passage, hESCs will take ∼7 days to reach ∼80% confluence. Each of the FACS procedures will take ∼4 hr from cell harvest to FACS analysis (longer if more flasks of cells are sorted). Preparation of total RNA takes ∼1 hr.

Acknowledgements Methods described in this unit were developed using funding from the Australian Stem Cell Centre (ASCC) to ALL. We would also like to thank the ASCC core hESC facility for provision of human embryonic stem cells, and the Flow Core [a collaborative initiative between Monash University, the ASCC, and the Australian Regenerative Medicine Institute (ARMI)] for FACS. We thank Martin Pera for the provision of GCTM-2 antibody.

Literature Cited Boyer, L.A., Lee, T.I., Cole, M.F., Johnstone, S.E., Levine, S.S., Zucker, J.P., Guenther, M.G., Kumar, R.M., Murray, H.L., Jenner, R.G., Gifford, D.K., Melton, D.A., Jaenisch, R., and Young, R.A. 2005. Core transcriptional regulatory circuitry in human embryonic stem cells. Cell 122:947-956. Chambers, I., Colby, D., Robertson, M., Nichols, J., Lee, S., Tweedie, S., and Smith, A. 2003. Func-

tional expression cloning of Nanog, a pluripotency sustaining factor in embryonic stem cells. Cell 113:643-655. Filipczyk, A.A., Laslett, A.L., Mummery, C., and Pera, M.F. 2007. Differentiation is coupled to changes in the cell cycle regulatory apparatus of human embryonic stem cells. Stem Cell Res. 1:45-60. Hough, S.R., Laslett, A.L., Grimmond, S.B., Kolle, G., and Pera, M.F. 2009. Metastable states of pluripotency in human embryonic stem cells. PLOS ONE 4:e7708. Kolle, G., Ho, M., Zhou, Q., Chy, H.S., Krishnan, K., Cloonan, N., Bertoncello, I., Laslett, A.L., and Grimmond, S.M. 2009. Identification of human embryonic stem cell surface markers by combined membrane-polysome translation state array analysis and immunotranscriptional profiling. Stem Cells 27:2446-2456. Laslett, A.L., Filipczyk, A., and Pera, M.F. 2003. Characterization and culture of human embryonic stem cells. Trends Cardiovasc. Med. 13:295-301. Laslett, A.L., Grimmond, S., Gardiner, B., Stamp, L., Lin, A., Hawes, S.M., Wormald, S., NikolicPaterson, D., Haylock, D., and Pera, M.F. 2007. Transcriptional analysis of early lineage commitment in human embryonic stem cells. BMC Dev. Biol. 7:12. Mitsui, K., Tokuzawa, Y., Itoh, H., Segawa, K., Murakami, M., Takahashi, K., Maruyama, M., Maeda, M., and Yamanaka, S. 2003. The homeoprotein Nanog is required for maintenance of pluripotency in mouse epiblast and ES cells. Cell 113:631-642. Pera, M.F., Filipczyk, A., Hawes, S.M., and Laslett, A.L. 2003. Isolation, characterization, and differentiation of human embryonic stem cells. Methods Enzymol. 365:429-446.

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SECTION 1C Culture and Maintenance of Undifferentiated Embryonic Stem Cells INTRODUCTION ll somatic and germ cells are derived from the inner cell mass (ICM) of blastocyst stage embryos. While the ICM is only a transient structure in vivo, stable in vitro cultures of pluripotential embryonic stem cells (ESCs) can be derived from it. How can ESCs be maintained in the undifferentiated state? Ever since the first derivation of ESCs from mice in 1981 it was clear that soluble factors are important for the expansion of ESCs. The differentiation inhibitory activity (DIA) critical for mouse ESCs was later shown to be leukemia inhibitory factor (LIF). Recently, the bone morphogenetic protein (BMP) pathway has emerged as a second major signaling pathway in mouse ESCs. Standard mouse ESC culture conditions consist of media containing LIF and fetal bovine serum (FBS; known to contain BMP-like activities) and use of gelatin-coated dishes or mouse embryonic fibroblasts (MEFs) as feeder cells.

A

With the recent derivation of primate and human ESCs it quickly became apparent that their optimal culture conditions differ significantly from those established for mouse ESCs: LIF is dispensable, and the BMP pathway should be inhibited rather than stimulated. Furthermore, addition of basic fibroblast growth factor (bFGF or FGF-2) and stimulation of the transforming growth factor beta pathway are critical. Accordingly, most laboratories do not use FCS in hESC media. Instead, a commercially available serum replacement, knockout serum replacement (KOSR; Invitrogen) containing albumin, insulin, transferrin, and other agents found in serum; is commonly used. Despite these differences, MEFs have also proven useful as feeder cells. There are several reasons for the development of chemically defined ESC growth conditions. (1) Conventional media exhibit significant lot-dependent variability that require extensive testing of undefined components such as FBS or KOSR and may compromise the reproducibility of results. (2) The ideal medium would only include components necessary for optimal cell growth and function, thereby simplifying the analysis of molecular mechanisms that regulate ESCs. (3) Exposure to undefined xeno-products (e.g., FBS, KOSR, or MEFs) poses significant risks of molecular or microbiological contamination that may limit the clinical utility of human ESCs and their derivatives. Chemically defined media that allow derivation and long-term expansion of mouse and human ESCs have recently been developed. An important difference between mouse and human ESC culture is that mouse ESCs can easily be passaged as single cells while human ESCs generally should be passaged as clumps. The reason for the more laborious and less efficient “clump passaging” of human ESCs is that long-term use of single-cell passaging techniques often leads to the emergence of karyotypically abnormal cells. However, techniques such as genetic manipulation, subcloning, and cell sorting are therefore significantly more challenging with clump-passaged human ESCs. Recovery of cryo-preserved human ESC clumps is also less efficient than that of frozen mouse ESCs. While the conventional method of slow freezing in the presence of DMSO yields acceptable results for at least shortterm storage of human ESC clumps, the more cumbersome vitrification technique is the

Current Protocols in Stem Cell Biology 1C.0.1-1C.0.2 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c00s1 C 2007 John Wiley & Sons, Inc. Copyright


Embryonic and Extraembryonic Stem Cells

1C.0.1 Supplement 1

current method of choice when both long-term storage and the highest rate of recovery are essential. In UNIT 1C.1, protocols for the expansion of karyotypically normal human ESCs are provided. The unit begins with a method for mechanically passaging human ESCs; this is followed by a method for enzymatic short-term bulk expansion. Thorsten M. Schlaeger

Introduction

1C.0.2 Supplement 1

Current Protocols in Stem Cell Biology

Expansion of Human Embryonic Stem Cells In Vitro

UNIT 1C.1

Magdaline Costa,1 Koula Sourris,1 Tanya Hatzistavrou,1 Andrew G. Elefanty,1 and Edouard G. Stanley1 1

Monash University, Clayton, Victoria

ABSTRACT This unit describes a protocol for the large-scale expansion of karyotypically normal human embryonic stem cells (hESCs). hESCs can be maintained indefinitely as dense colonies that are mechanically cut into pieces, which are subsequently transferred to fresh organ culture dishes seeded with primary mouse embryonic fibroblasts (MEFs). hESCs can also be enzymatically passaged (bulk culture); however, over time, this style of culturing may lead to the acquisition of chromosomal abnormalities. Nevertheless, enzymatic passaging can be used for short periods (up to 25 passages) without the appearance of cells with abnormal karyotypes. Curr. Protoc. Stem Cell Biol. 5:1C.1.1C 2008 by John Wiley & Sons, Inc. 1C.1.7.
Keywords: human embryonic stem cells (hESCs) r mechanical passaging r enzymatic passaging r hESC expansion

INTRODUCTION This unit describes a protocol for the large-scale expansion of karyotypically normal human embryonic stem cells (hESCs). hESCs can be maintained indefinitely as dense colonies that are mechanically cut into pieces, which are subsequently transferred to fresh organ culture dishes seeded with primary mouse embryonic fibroblasts (MEFs; Thomson et al., 1998; Reubinoff et al., 2000). hESCs can also be enzymatically passaged (bulk culture; Amit et al., 2000); however, over time, this style of culturing may lead to the acquisition of chromosomal abnormalities (Draper et al., 2004). Nevertheless, enzymatic passaging can be used for short periods (up to 25 passages) without the appearance of cells with an abnormal karyotype or changes detected by morphological and FACS analysis. This unit begins with a method for the propagation of hESCs in organ culture dishes (Basic Protocol 1), followed by a protocol for large-scale expansion of hESCs (Basic Protocol 2) that can be used subsequently for experiments. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

MAINTENANCE OF hESC CULTURES BY MECHANICAL PASSAGING (MAINTENANCE CULTURE) This protocol is used for the long-term maintenance of hESCs. During culture in organ culture dishes, the cells are nonenzymatically passaged and can be continuously cultured for over 2 years without the acquisition of an abnormal karyotype. Cells are maintained Current Protocols in Stem Cell Biology 1C.1.1-1C.1.7 Published online April 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c01s5 C 2008 John Wiley & Sons, Inc. Copyright


BASIC PROTOCOL 1

Embryonic and Extraembryonic Stem Cells

1C.1.1 Supplement 5

Expansion of hESC In Vitro

Figure 1C.1.1 Photomicrographs of hESCs during various stages of culture. (A) Overview of day 5 hESC colonies grown on MEFs. (B) and (C) 50× magnification of a single colony under phase-contrast and bright-field visualization, respectively. The colony has a dense raised area. (D) Overview of day 5 hESC colonies grown on MEFs with the central raised areas eliminated. (E) and (F) 50× magnification of a single colony under phase contrast and bright field, respectively, with the central raised area eliminated. (G) and (H) 50× magnification of the same colony shown in (E) and (F) 2 days later (day 7) under phase contrast and bright field, respectively. (I) and (J) 50× magnification of a day 7 colony under phase contrast and bright field, respectively, sliced into a grid motif prior to dislodgement and transfer to new organ culture dishes. (K) 50× magnification of hESCs in bulk culture passage 1. (L) 50× magnification of hESC in bulk culture passage 2 after treatment with TrypLE Select. (M) 50× magnification of hESCs in bulk culture on feeders at reduced density on day of application.

1C.1.2 Supplement 5

Current Protocols in Stem Cell Biology

in organ culture dishes as dense colonies and passaged once every 7 days based on published methods (Thomson et al., 1998; Reubinoff et al., 2000) with modifications as described.

Materials hESCs, starting from macroscopic colonies (∼1-mm diameter) grown on MEFs (UNIT 1A.2) Mitotically inactivated (irradiation- or mitomycin C–treated; UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000; Nagy et al., 2003) hESC medium (see recipe) Stereomicroscope 26-G × 1/2 -in. (0.45 × 13–mm) needles 1-ml syringe 60 × 15–mm center-well organ culture dishes Additional reagents and equipment for hESCs grown on feeder cells (UNIT 1A.2) and mitotically inactivated mouse embryonic fibroblasts (Conner, 2000; UNIT 1C.3) Maintain hESCs in organ culture dishes 1. At a time point 2 days before transfer, excise differentiated and/or raised regions within each colony using a 26-G needle attached to a 1-ml syringe (Fig. 1C.1.1, A through F). These procedures are performed under a stereomicroscope.

2. On the day before transfer, plate mitotically inactivated MEFs (Conner, 2000; UNIT 1C.3) onto the center well of gelatinized organ culture dishes at a density of 6 × 104 cells/cm2 in 1 ml MEF medium. 3. On the day of transfer, day 7, using a 26-G needle, cut each colony into a grid motif containing approximately eight pieces per colony and dislodge with the same 26-G needle (Fig. 1C.1.1, G through J). 4. Replace medium in organ culture dishes (prepared in step 2) with 1 ml hESC medium and transfer up to ten pieces into the center well. 5. Change the medium to fresh hESC medium daily. Using this method a single organ culture plate containing 10 colonies should yield ∼18 organ culture plates after 3 weeks of passaging—a sufficient number to enter the enzymatic passaging protocol described below.

EXPANSION OF hESC IN BULK CULTURE This protocol is used for the large-scale expansion of hESCs. In the authors’ experience, the cells can be enzymatically passaged during this bulk culture stage up to 25 times without the acquisition of an abnormal karyotype if the following procedure is followed. It is recommended that bulk culture begin with the lowest-passage-number cells available. The authors have also found that, if it is necessary to preserve a valuable reagent, returning genetically modified, enzymatically passaged cells to maintenance culture results in cultures that appear indistinguishable (morphology, FACS phenotype, karyotype, gene expression) from parental cultures.

BASIC PROTOCOL 2

Materials hESCs in organ culture dishes (Basic Protocol 1) hESC medium (see recipe) Trypsin (see recipe) or TrypLE Select (Invitrogen) Phosphate-buffered saline without CaCl2 , without MgCl2 (CMF-PBS) Soybean Trypsin Inhibitor (Invitrogen), optional Current Protocols in Stem Cell Biology

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26-G × 1/2 -in. (0.45 × 13–mm) needles 1-ml syringe 200-µl Gilson pipet (optional) 75-cm2 flask containing preseeded mitotically inactivated MEFs at a density of 4 × 104 cells/cm2 , 2 × 104 cells/cm2 , and 1 × 104 cells/cm2 (Connor, 2000; UNIT 1C.3) 150-cm2 tissue culture flask with vented cap Stereomicroscope Additional reagents and equipment for preparing mitotically inactivated mouse embryonic fibroblasts (Connor, 2000; UNIT 1C.3), growing hESCs in organ culture dishes (Basic Protocol 1), cell counting (Phelan, 2006; UNIT 1C.3), and electroporation (Costa et al., 2007) Expand ES cells in bulk culture 1. Using a 26-G needle attached to a 1-ml syringe, cut colonies from 18 organ culture dishes (∼10 colonies/dish) into a grid motif to generate ∼25 small pieces per colony. 2. Dislodge the sliced colonies from the dish either with the same needle or with a 200–µl Gilson pipet. Collect these small pieces into a 15-ml centrifuge tube. 3. Centrifuge the pieces 3 min at 480 × g, 4◦ C. Resuspend the entire pellet in 10 ml hESC medium and plate onto a gelatinized 75-cm2 flask containing irradiated MEFs seeded previously at a density of 4 × 104 cells/cm2 (Fig. 1C.1.1K). This is designated as the first passage in bulk culture. At this stage the cells have not been exposed to any enzyme and are still tightly bound together in small clumps.

4. At a time point 3 days later, wash the hESCs once with 3 ml CMF-PBS. The authors have found that enzymatically passaging smaller hESC colonies at 3 days results in a higher survival rate than passaging larger colonies, which are more sensitive to trypsinization, at 5 to 7 days.

5. Add 2 ml of trypsin or TrypLE Select and incubate for 5 min at 37◦ C or until the cells have dislodged from the dishes. If trypsin is used as the dissociation agent, then a neutralization step is required such as washing the cells with 10 ml serum-containing medium or adding 1 ml soybean trypsin inhibitor.

6. During these first enzymatic passages, break up hESC colonies by trituration to produce predominantly single cells with some small clumps remaining. 7. After trypsinization, collect the cells by centrifuging 3 min at 480 × g, 4◦ C. 8. Resuspend the cells in 10 ml hESC medium and transfer to a fresh gelatinized 75-cm2 flask preseeded the day before with MEFs at a density of 2 × 104 cells/cm2 (Fig. 1C.1.1L). In the authors’ experience, there is usually extensive cell death associated with this first enzymatic passage (50%). The amount of cell death decreases with subsequent passages.

9. Passage the cells enzymatically twice a week (refer to steps 4 to 8), to ensure that the colonies remain small. This generally allows the cultures to be expanded ∼1:2 on each passage. For the firsttime passaging to 150-cm2 flasks, transfer the contents of a 75-cm2 flask to one 150-cm2 flask (1:2 passage). Expansion of hESC In Vitro

By this stage the cell numbers increase rapidly and by passage 4 to 5 there are enough cells for applications such as electroporation (Costa et al., 2007), transfection, or differentiation.

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Day before application 10. Enzymatically passage cells (refer to steps 4 to 8) and transfer the cells into gelatinized 150-cm2 flasks containing irradiated MEFs seeded at a density of 1 × 104 cells/cm2 . A lower MEF density is required for enzymatically passaged hESCs than for those in maintenance culture. This may be related, in part, to the fact that the MEFs are replaced more often as a consequence of more frequent passaging. As a rule, on the day of application, aim for a semiconfluent flask of cells, i.e., ∼8 × 106 hESC/150 cm2 .

Day of application 11. In the morning, change the medium on the cells to fresh hESC medium (Fig. 1C.1.1M). 12. Harvest cells using 2 ml trypsin or TrypLE Select (steps 4 to 5) after first washing with CMF-PBS. If trypsin is used as the dissociation agent, then a neutralization step is required such as washing the cells with 10 ml serum-containing medium or adding 1 ml soybean trypsin inhibitor.

13. Perform a viable cell count (Phelan, 2006; UNIT 1C.3) and subtract total feeder number from the count for an estimate of the number of hESCs harvested. For example, an area of 150-cm2 will contain ∼1.5 × 106 feeders in the total cell count.

14. Use the cells for electroporation (Costa et al., 2007), transformation, or differentiation.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Gelatin 0.1% (w/v) Dissolve 0.5 g of gelatin (from porcine skin) in 500 ml distilled water and autoclave. Store at room temperature indefinitely.

Gelatinized flasks/plates Prior to addition of MEFs, coat all plates and flasks with enough 0.1% (w/v) gelatin solution (see recipe) to cover the surface. Remove gelatin after 5 min.

hESC medium DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (Invitrogen) 10 mM non-essential amino acids 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) 50 mM 2-mercaptoethanol 10 ng/ml bFGF (Amit et al., 2000; see recipe) Store up to 1 week at 4◦ C MEF medium DMEM containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C

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Recombinant human basic fibroblast growth factor (bFGF) Resuspend lyophilized bFGF (PeproTech) to a final concentration of 10 µg/ml in CMF-PBS containing 0.1% (w/v) BSA and 1 mM DTT. Store at –80◦ C according to manufacturer’s instructions.

Trypsin, 0.125% (w/v) Trypsin/EDTA (0.25% trypsin EDTA; Invitrogen, no. 25200-056) supplemented with:

2% (v/v) chicken serum (Hunter Antisera, no. 110) Dilute 1:2 with CMF-PBS Store for 4 weeks at 4◦ C COMMENTARY Background Information

Expansion of hESC In Vitro

ESCs are pluripotent cells that are isolated from the inner cell mass of the blastocyststage embryo and can be cultured indefinitely in vitro (Evans and Kaufman, 1981; Martin, 1981; Thomson et al., 1998). Their ability to differentiate into multiple cell types (Evans and Kaufman, 1981; Nagy et al., 1993; Thomson et al., 1998; Reubinoff et al., 2000) makes them a suitable substrate for studies involving drug discovery (McNeish, 2004), human development, and cell therapies (Menendez et al., 2005). hESCs are often cultured as dense colonies which are mechanically cut into small pieces and transferred from one organ culture dish to another (Thomson et al., 1998; Reubinoff et al., 2000). This method is preferred for longterm maintenance of hESCs as it reduces the level and frequency of stress associated with passaging and reduces the incidence of karyotypic abnormalities. Collagenase Type IV is commonly used for the large-scale passaging of hESCs (Amit et al., 2000); however, because this enzyme usually yields cell clumps, precise cell numbers can only be estimated making this method unsuitable for situations where precise cell numbers or single cells are required. Replacing Collagenase Type IV with either trypsin or TrypLE Select enables hESCs to be passaged as a single-cell suspension. All enzymatic passaging methods eventually select for cells which are adapted to such methods. Over time, such adaptations may include the acquisition of chromosomal aberrations (Draper et al., 2004) that provide a selective advantage to cells grown under these conditions. For this reason it is recommended that enzymatic passaging only be used transiently for the generation of large cell numbers required for experiments rather than as a method for routine long-term hESC maintenance. Uses

for enzymatically passaged cells include electroporation of ∼1 × 107 cells, typical spin EB (embryoid bodies) method for differentiation of 3 × 105 hESCs per 96-well plate with 3000 cells per well. Most experiments would use 10 to 30 plates.

Critical Parameters and Troubleshooting hESCs are cultured using two different techniques, each with different requirements. The two techniques are listed below. Maintenance culture in organ culture dishes: the feeder density influences the thickness of the hESC colony. If the feeder density is too high then the colonies will be very thick and will tend to tear when being cut for passaging. If the feeder density is sparse, the colonies will be thin and the cut pieces will fray when being manually dislodged from the dish during passaging. Bulk culture: The time it takes to achieve confluence from one passage to the next is influenced by the number and size of colonies present in the initiating culture. Although the hESCs are eventually passaged as a single-cell suspension they reform colonies on the dish as the cells proliferate. The longer these colonies are left to regrow between passages, the harder it is to dissociate them, which in turn leads to higher levels of cell death once passaged. Passaging the cells twice a week regardless of the number of colonies per flask prevents the colonies from becoming too large and difficult to dissociate.

Anticipated Results This protocol generates large numbers of karyotypically normal hESCs, suitable for numerous applications such as electroporation and differentiation. After 5 bulk (enzymatic)

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passages, expect 4 × 150-cm2 flasks each containing ∼8 × 106 hESCs (∼3.2 × 107 total).

Time Considerations hESC colonies from one organ culture dish containing 10 colonies should be able to be distributed among six new dishes. It should take 2 weeks to go from 1 dish to 18. During the third week the colonies on the 18 dishes are mechanically transferred to a single 75cm2 flask (passage 1) which is subsequently passaged 3 days later (passage 2) using either trypsin or TrypLE select. By passage 4 (week 4), there should be sufficient cells to generate two confluent 150-cm2 flasks. At this stage, cells which are to be used for experiments are passaged onto flasks seeded with MEFS at a reduced density (passage 5). Alternatively, cells can be enzymatically passaged 20 to 25 times without the appearance of chromosomal abnormalities. Under such circumstances, excess cells generated at each passage can be fed into other applications, kept for future experiments, or discarded.

Acknowledgement We thank Elizabeth Ng for her valuable contribution to the development of the maintenance culture protocol described in this unit.

Literature Cited

S., Lim, S-M., Pera, M., Elefanty, A.G., and Stanley, E.G. 2007. A method for genetic modification of human embryonic stem cells using electroporation. Nat. Protoc. 2:792-796. Draper, J.S., Smith, K., Gokhale, P., Moore, H.D., Maltby, E., Johnson, J., Meisner, L., Zwaka, T.P., Thomson, J.A., and Andrews, P.W. 2004. Recurrent gain of chromosomes 17q and 12 in cultured human embryonic stem cells. Nat. Biotechnol. 22:53-54. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Martin, G.R. 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. 78:76347638. McNeish, J. 2004. Embryonic stem cells in drug discovery. Nat. Rev. Drug Discov. 3:70-80. Menendez, P., Wang, L., and Bhatia, M. 2005. Genetic manipulation of human embryonic stem cells: A system to study early human development and potential therapeutic applications. Curr. Gene Ther. 5:375-385. Nagy, A., Rossant, J., Nagy, R., AbramowNewerly, W., and Roder, J.C. 1993. Derivation of completely cell culture-derived mice from early-passage embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 90:8424-8428. Nagy, A., Gertsenstein, M., and Vintersten, K. 2003. Manipulating the Mouse Embryo: A Laboratory Manual, 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.

Amit, M., Carpenter, M.K., Inokuma, M.S., Chiu, C.P., Harris, C.P., Waknitz, M.A., ItskovitzEldor, J., and Thomson, J.A. 2000. Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227:271278.

Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A3F.1-A.3F.18.

Conner, D.A. 2000. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Mol. Biol. 51:23.2.1-23.2.7.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

Costa, M., Dottori, M., Sourris, K., Jamshidi, P., Hatzistavrou, T., Davis, R., Azzola, L., Jackson,

Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399-404.

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Defined, Feeder-Independent Medium for Human Embryonic Stem Cell Culture

UNIT 1C.2

Tenneille Ludwig1,2 and James A. Thomson1,2,3 1

University of Wisconsin-Madison, Madison, Wisconsin WiCell Research Institute, Madison, Wisconsin 3 University of Wisconsin School of Medicine and Public Health, Madison, Wisconsin 2

ABSTRACT The developmental potential of human ES cells makes them an important tool in developmental, pharmacological, and clinical research. For human ES cell technology to be fully exploited, however, culture efficiency must be improved, large-scale culture enabled, and safety ensured. Traditional human ES cell culture systems have relied on serum products and mouse feeder layers, which limit the scale, present biological variability, and expose the cells to potential contaminants. Defined, feeder-independent culture systems improve the safety and efficiency of ES cell technology, enabling translational research. The protocols herein are designed with the standard research laboratory in mind. They contain recipes for the formulation of mTeSR (a defined medium for human ES cell culture) and detailed protocols for the culture, transfer, and passage of cells grown in these feederindependent conditions. They provide a basis for routine feeder-independent culture, and a starting point for additional optimization of culture conditions. Curr. Protoc. Stem Cell C 2007 by John Wiley & Sons, Inc. Biol. 2:1C.2.1-1C.2.16.
Keywords: feeder-independent culture r human ES cells r defined medium r bFGF

INTRODUCTION The developmental potential of human ES cells means that they have tremendous potential to be a useful tool in elucidating the early stages of development, advancing pharmacological research, and improving human health. For human ES cell technology to be fully exploited, however, culture efficiency must be improved and large-scale culture enabled. Translating human ES cell technology to clinical applications will also make safety of paramount importance. Traditional human ES cell culture systems rely on poorly defined serum products and mouse embryonic feeder layers (MEFs). The inclusion of these ill-defined components in the culture system significantly reduces the efficiency of human ES cell culture and exposes the cells to potential contaminants from animal-sourced proteins. Development and refinement of defined culture systems that eliminate the need for feeder layers, while maintaining undifferentiated proliferation, will improve the safety and efficiency of human EC cell culture and enable translational research. The protocols in this unit are designed with the standard research laboratory in mind, and contain complete recipes for the formulation of mTeSR (see Reagents and Solutions), a defined medium for the feeder-independent propagation of human ES cells. Also included are detailed protocols for the transfer and culture (see Basic Protocol 1), and passage (see Basic Protocol 2, Alternate Protocols 1 and 2) of cells grown in these feederindependent conditions. These recipes and protocols are intended to provide a basis for routine feeder-independent culture, and a starting point for additional optimization of culture conditions. Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1C.2.1-1C.2.16 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c02s2 C 2007 John Wiley & Sons, Inc. Copyright


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NOTE: For all procedures described in this unit, standard tissue culture, reagent preparation, and sterilization facilities are required. All cell handling should be performed under sterile conditions in a Class II Biological Hazard Flow Hood, and all biologically contaminated material should be disposed of properly. NOTE: All cell cultures should be maintained at 37◦ C in a humidified atmosphere of 5% CO2 in air. NOTE: The use of human embryonic stem cells as described in these protocols usually requires specific MTA approval from the appropriate institutional research office, and may require ethics approval from the appropriate institutional committee. NOTE: The authors recommend glass serological pipets for all measured media transfers, unless specifically indicated otherwise. BASIC PROTOCOL 1

TRANSFERRING HUMAN EMBRYONIC STEM CELLS TO AND CULTURING IN FEEDER-INDEPENDENT CONDITIONS The move to defined, feeder-independent culture systems can significantly increase the overall efficiency of the research laboratory. First and foremost, it eliminates the need for the labor-intensive derivation, maintenance, and routine preparation of MEFs otherwise necessary for culture or conditioning of media. Furthermore, the removal of highly variable components (including serum and MEFs) from the culture system results in more consistent, reliable, repeatable results, speeding the progress of research. The basic protocols for the feeder-independent culture of human ES cells outlined below are modifications of those originally published in Ludwig et al. (2006a).

Materials Human ES cells in standard (MEF or feeder-free) culture, in 6-well plates mTeSR culture medium (see recipe) Matrigel-coated 6-well plates (see recipe) Transfer to feeder-independent conditions 1. At a time point 3 days prior to normal passage time, replace current culture medium with 2 ml/well warmed mTeSR culture medium. While the authors recommend only warming aliquots of the medium, all of their testing was done while repeatedly warming the entire bottle. Since this is the method employed by the general scientific community, they wanted to ensure that the medium would perform under these circumstances.

2. Feed cultures daily with 2 ml mTeSR medium until ready to passage. Spent culture medium should be completely removed at each feeding and replaced with 2 ml/well warmed (37◦ C) mTeSR culture medium.

3. Passage cells using either Alternate Protocol 1 or Alternate Protocol 2, as appropriate. Plate cells directly onto Matrigel-coated 6-well plates with 2 ml of mTeSR medium at a density of ∼5 × 105 per well. On occasion, the authors have used both 35- and 60-mm dishes. This protocol is designed for 6-well plates, but could easily be adapted by adjusting media volumes to an alternate format. Cells should be passaged more densely than normal for this transfer passage. Defined, FeederIndependent Medium for hESC Culture

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If passaging from feeder-free (conditioned medium) cultures on Matrigel, the authors recommend transfer passaging as outlined in Alternate Protocol 1. If passaging from MEF-containing cultures, the authors recommend transfer passaging as outlined in Alternate Protocol 2. While the authors routinely transfer MEF cultures to mTeSR culture systems, some users have preferred first establishing cultures in a MEF-conditioned medium system before moving to feeder-independent culture systems.

Feeder-independent culture 4. Feed and examine cultures daily until ready to passage (∼5 days). Each day, aspirate all spent medium and replace with 2 ml/well warmed (37◦ C) mTeSR culture medium. Medium volume/well is based on a single well of a standard 6-well plate with a surface area of 9.6 cm2 . If alternate size surface area is being used, adjust medium volume accordingly.

PASSAGING HUMAN EMBRYONIC STEM CELLS IN FEEDER-INDEPENDENT CONDITIONS

BASIC PROTOCOL 2

There is some evidence to suggest that manual passaging may result in more stable cultures than those enzymatic techniques that individualize cells at passage (Mitalipova et al., 2005). For this reason it has been recommended to reduce the accumulation of karyotypic abnormalities (Buzzard et al., 2004; Mitalipova et al., 2005). While manual passaging results in more consistent clump size at passage, it is labor intensive and incompatible with large-scale culture systems necessary for the industrial and clinical use of human ES cells. Alternatives to manual passaging include those bulk passaging systems (both enzymatic and non-enzymatic) that allow cultures to be primarily passaged in clumps, with limited individualization of cells. If appropriate care is taken to ensure that colonies are passaged on time and are not excessively disrupted, these bulk passaging methods described below can be used routinely with success. The authors have cultured cells for >100 passages using enzymatic passaging and maintained a normal karyotype, and routinely propagate cultures for >30 passages using all of these systems with no adverse effect.

Materials Human ES cell culture in Matrigel coated 6-well plates (Basic Protocol 1) mTeSR culture medium (see recipe) Washing medium (see recipe) EDTA splitting medium (see recipe) Inverted microscope with marking objective (Nikon) Pasteur pipets (Fisher Scientific) 15-ml conical tube (optional) Glass serological pipets (Fisher Scientific) Prepare Matrigel plate 1. Prepare Matrigel plate for use by aspirating excess Matrigel and plating 2 ml/well of mTeSR culture medium into each well. Label plate appropriately, and set aside. 2. Observe cultures using phase contrast microscopy. Mark any small area of differentiation to be removed prior to passage with the marking objective (Fig. 1C.2.1). 3. Using a Pasteur pipet, aspirate spent medium. During aspiration, touch pipet to marked area to remove differentiating cells (Fig. 1C.2.1). 4. Wash culture twice with 1 ml/well of washing medium. 5. Aspirate washing medium and replace with 1 ml/well EDTA splitting medium.

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Figure 1C.2.1 Removal of areas of differentiation by aspiration. Using phase contrast microscopy, view cultures prior to passage (A) If small areas of differentiation are noted, they should be marked using a marking objective. (B) After aspirating medium, touch the Pasteur pipet to the marked area to remove differentiated area. (C) Scale bar = 100 µm.

6. Incubate 1 to 2 min at room temperature. Be cautious not to over-incubate, as cells will detach prematurely. Incubation for >3 min will make it impossible to split the cells without centrifugation.

7. Aspirate splitting medium and replace immediately with mTeSR culture medium. 8. Using a serological pipet, remove colonies from the plate by gently pipetting the medium against the bottom of the plate, releasing the colonies. In general, scraping is not necessary, but if cells do not dislodge immediately, use the serological pipet and gently scrape the bottom while simultaneously expelling medium.

9. If multiple wells are being passaged, pool the cell suspensions into a 15-ml conical tube. 10. Gently mix by pipetting the colonies to ensure even distribution, being careful not to disrupt the colonies more than necessary. Average colony size should be no smaller than 50 to 100 cells.

Passage cells 11. Passage cells so that roughly 2 × 105 cells are seeded into each fresh well. Generally this translates into a 1:8 to 1:15 split every 7 days.

12. Gently shake the plate to evenly distribute the colonies, and return to the incubator.

Culture cells 13. Culture as per Basic Protocol 1 until cells are again ready to passage.

Defined, FeederIndependent Medium for hESC Culture

Cultures must be passaged at the appropriate time to ensure continued quality. While other culture methods allow for some flexibility in split timing, in this feeder-independent system there is only a 12 to 24 hr window in which to passage cells and achieve optimum attachment and continued undifferentiated proliferation. Cells will easily differentiate if allowed to overgrow, and cannot be rescued. Passaging too early, however, results in poor attachment and limited growth. Cultures should be passaged when colony centers become dense, appearing brighter than the edges when viewed using phase contrast microscopy (Fig. 1C.2.2). Some users may find it helpful to split sister wells of a single culture on successive days when initially working with the system. Observation of the individual cultures in the days immediately following passage may assist in accurately identifying the appropriate passage time.

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Figure 1C.2.2 Identifying appropriate passage timing. Cultures should be passaged just as the centers of the colonies become dense, appearing brighter than the edges using phasecontrast microscopy. If split too early, attachment will be reduced. If allowed to overgrow, cells will differentiate, and cannot be rescued. Scale bar = 100 µm.

PASSAGING WITH DISPASE In addition to EDTA (Basic Protocol 2), dispase may also be used to enzymatically passage cells in bulk while maintaining appropriate karyotypes. The authors have done extensive screening in their laboratory, and in their hands, and while collagenase works best for passaging cells in co-culture, they achieve the best results using dispase or EDTA for passaging cells on Matrigel. Better initial attachment and expansion was seen using these methods. The dispase-based technique may also be used to routinely passage feeder-independent cultures, although scale-up can be achieved more rapidly using Basic Protocol 2.

ALTERNATE PROTOCOL 1

Materials Human ES cell culture in Matrigel-coated 6-well plates (for transfer or from Basic Protocol 1) mTeSR culture medium (see recipe) Dispase splitting medium (see recipe) Warmed DMEM/F-12 (Invitrogen) Inverted microscope with marking objective (Nikon) Pasteur pipets (Fisher Scientific) 37◦ C incubator 15-ml conical tube (optional) Glass serological pipets (Fisher Scientific)

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Prepare Matrigel plate and remove cells 1. Prepare Matrigel plate for use by aspirating excess Matrigel and plating 2 ml/well of mTeSR culture medium into each well. Label plate appropriately, and set aside. 2. Observe cultures using phase contrast microscopy. Mark any small area of differentiation to be removed prior to passage with the marking objective (Fig.1C.2.1). 3. Using a Pasteur pipet, aspirate spent medium. During aspiration, touch pipet to marked area to remove differentiating cells (Fig. 1C.2.1). 4. Add 1 ml/well dispase splitting medium and incubate plate 7 min at 37◦ C. 5. Following incubation, aspirate splitting medium and gently rinse cells on the plate a minimum of three times using 1 ml warmed DMEM/F-12 medium. Adequate rinsing at this step is critical, and reducing the number of washes will dramatically reduce or prevent colony plating.

6. Aspirate rinse medium and gently remove colonies from the plate by rinsing with 1 to 2 ml mTeSR culture medium, gently scraping the bottom of the plate as necessary. In general, scraping is not necessary, but if cells do not dislodge immediately, use the serological pipet and gently scrape the bottom while simultaneously expelling medium.

7. If multiple wells are being passaged, pool the cell suspensions into a 15-ml conical tube. 8. Gently mix by pipetting the colonies to ensure even distribution, being careful not to disrupt the colonies more than necessary.

Passage cells 9. Passage cells so that roughly 3 × 105 cells are seeded into each fresh well. This generally translates into a 1:3 to 1:6 split every 4 to 5 days.

10. Gently shake the plate to evenly distribute the colonies, and return to the incubator.

Culture cells 11. Culture as per Basic Protocol 1 until cells are again ready to passage. ALTERNATE PROTOCOL 2

PASSAGING WITH COLLAGENASE The best results the authors have obtained to date with enzymatically passaging human ES cell cultures on MEFs have been achieved using collagenase. The authors do not recommend this technique for passaging human ES cells grown on Matrigel or other extracellular matrices: it is used exclusively in their laboratory for the passaging of human ES cell cultures in direct contact with MEFs. The authors recommend this technique when transferring cultures from MEFs to feeder-independent culture systems. Once cultures have been established in feeder-independent systems, the authors recommend one of the above protocols (Basic Protocol 2 or Alternate Protocol 1) for continued passaging.

Materials Human ES cell culture on MEFs (for transfer to feeder-independent systems) mTeSR culture medium (see recipe) Collagenase splitting medium (see recipe) Warmed DMEM/F-12 (Invitrogen) Defined, FeederIndependent Medium for hESC Culture

Inverted microscope with marking objective Pasteur pipets (Fisher Scientific) 15-ml centrifuge tube (optional) Glass serological pipets (Fisher Scientific)

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Prepare Matrigel plate and remove cells 1. Prepare Matrigel plate for use by aspirating excess Matrigel and plating 2 ml/well of mTeSR culture medium into each well. Label plate appropriately, and set aside. 2. Observe cultures using phase contrast microscopy. Mark any small area of differentiation to be removed prior to passage with the marking objective (Fig. 1C.2.1). 3. Using a Pasteur pipet, aspirate spent medium. During aspiration, touch pipet to marked area to remove differentiation (Fig. 1C.2.1). 4. Add 1 ml/well collagenase splitting medium and incubate plate at 37◦ C for 5 min. To confirm colony separation from the plate, view surface under a microscope. Look for the perimeter of the colony to appear folded back. If necessary, keep collagenase on cells for another minute or two. 5. Following incubation, aspirate splitting medium and gently rinse cells on the plate using 1 ml/well warmed DMEM/F-12 medium. 6. Aspirate rinse medium and gently remove colonies from the plate by rinsing with 1 to 2 ml mTeSR culture medium. In general, scraping is not necessary, but if cells do not dislodge immediately, use the serological pipet and gently scrape the bottom while simultaneously expelling medium.

7. If multiple wells are being passaged, pool the cell suspensions into a 15-ml conical tube. 8. Using a serological pipet, break up colonies by gently pipetting up and down a few times, and gently mix the colonies to ensure even distribution.

Passage cells 9. Passage cells so that roughly 3 × 105 cells are seeded into each fresh well. This generally translates into a 1:3 to 1:6 split every 4 to 5 days.

10. Gently shake the plate to evenly distribute the colonies, and return to the incubator.

Culture cells 11. Culture as per Basic Protocol 1 until cells are again ready to passage. REAGENTS AND SOLUTIONS For suppliers, see SUPPLIERS APPENDIX. Whenever possible, cell culture–tested reagents have been used, and are recommended by the authors. Water quality is critical, and only Type 1 reagent-grade water should be used in the recipes and protocols presented here. Good quality, Type 1 reagent-grade water can be obtained from a Milli-Q Ultrapure Water System (Millipore), and is best used immediately after drawing. Alternatively, if reasonable in-house purification systems are not available, Type 1 reagent-grade water is available for purchase from a few select vendors.

bFGF stock (50 µg/ml) Reconstitute 500 µg bFGF [either zebrafish (Ludwig et al., 2006a) or human (Peprotech)] in 10 ml diluent solution (see recipe). Use immediately or freeze in 10-ml aliquots for up to 6 months at −80◦ C.

Embryonic and Extraembryonic Stem Cells

1C.2.7 Current Protocols in Stem Cell Biology

Supplement 2

Collagenase splitting medium (1 mg/ml) Add 25 mg collagenase IV (Sigma) to 25 ml DMEM/F-12 medium (Invitrogen). Mix until dissolved. Filter sterilize the resulting medium and store up to 2 weeks at 4◦ C. Warm aliquot to 37◦ C before use.

Diluent solution (0.1% w/v BSA in PBS) Dissolve 100 mg BSA (Sigma) in 100 ml Ca/Mg-free PBS (Invitrogen). Filter sterilize the resulting solution, and store in 10-ml aliquots up to 6 months at 4◦ C.

Dispase splitting medium (2 mg/ml) Add 50 mg dispase (Sigma) to 25 ml DMEM/F-12 medium (Invitrogen). Mix until dissolved. Filter sterilize the resulting medium and store up to 2 weeks at 4◦ C. Warm an aliquot to 37◦ C before use.

EDTA splitting medium Add 1.48 g calcium- and magnesium-free DMEM/F-12 powder (Invitrogen) and 15 mg EDTA acid (anhydrous, crystalline, cell culture tested; Sigma) to 100 ml Type I water. Stir until dissolved, adding low heat as necessary. Once dissolved, adjust the pH of the medium to 7.2 and adjust the osmolarity to 340 ±5 mOsM using sodium chloride (Sigma). Filter sterilize and store up to 2 weeks at 4◦ C. Warm aliquot to 37◦ C before use. As the DMEM/F-12 base medium is currently only available in quantities sufficient to make 10-liter volumes, the authors recommend that it be distributed into 1.48 g aliquots for ease of use. These aliquots should be stored desiccated at −80◦ C for no more than 6 months.

L-glutamine solution Dissolve 146 mg L-glutamine (Sigma) into 10 ml Type 1 water. Add 7 µl-2mercaptoethanol solution (Sigma). Use immediately.

Matrigel-coated plates Thaw 0.5 mg (one vial) Matrigel from Matrigel stock (see recipe) by diluting into 6 ml of cold DMEM/F-12 (this should take no more than a minute). Mix well, and plate 1 ml of the resulting solution into each well of a 6-well culture plate. Allow Matrigel to settle at room temperature for at least 1 hr before use. Aspirate excess Matrigel from the plate immediately before use (plates need not be rinsed). Plates not used on the day plated can be wrapped with Parafilm or foil and stored at 4◦ C for up to 1 week. Warm plates to room temperature before use. The authors have noted significant reductions in plating efficiency if the Matrigel solution is allowed to dry prior to plating cells. Therefore, discard any stored plates that have dried prior to use.

Matrigel stock Thaw one bottle of Growth Factor Reduced (GFR) Matrigel (Becton Dickinson) on ice overnight at 4◦ C. Keeping the Matrigel on ice at all times and using chilled tips, aliquot 0.5 mg of Matrigel into prelabeled, prefrozen (−80◦ C) 1.5-ml tubes on ice. Immediately freeze tubes at −80◦ C.

Defined, FeederIndependent Medium for hESC Culture

The concentration of Matrigel varies from lot to lot, so the volume of Matrigel needed to obtain 0.5 mg will vary accordingly. Aliquoted as above, each tube will yield one coated 6-well plate. If multiple plates are desired, increase the volume of the aliquot accordingly. Do not allow Matrigel to warm at any point during this procedure. Doing so will cause the product to gel prior to use, resulting in uneven plating and reduced performance.

1C.2.8 Supplement 2

Current Protocols in Stem Cell Biology

mTeSR Culture medium Combine 800 ml DMEM/F-12 (Invitrogen) with 200 ml stock B (see recipe). Supplement with:

1% (v/v) non-essential amino acids (Invitrogen) 1% (v/v) L-glutamine solution (see recipe) Adjust pH to 7.4 using 10 N sodium hydroxide (Sigma) and adjust the osmolarity to 340 ±5 mOsM using sodium chloride (Sigma). Filter sterilize the resulting medium (mTeSR) and store for up to 2 weeks at 4◦ C. If culture medium has been prepared using fresh (not frozen/thawed) stock B, then whole medium may be frozen in aliquots and stored up to 6 months at −80◦ C. Warm aliquot to 37◦ C before use. There is no significant impact on culture performance when using medium that has been through a single freeze/thaw cycle (Fig. 1C.2.3).

Pipecolic acid stock (100 mg/ml) Dissolve 1 g L-pipecolic acid (MP Biomedicals) in 10 ml diluent solution (see recipe). Store 1- to 2-ml aliquots up to 6 months at −80◦ C.

Figure 1C.2.3 Effect of freezing medium on cell competence. Human ES cells were cultured in either fresh or lot-matched frozen medium for three passages. At the end of the culture period, Oct4 expression was assessed by FACS analysis (B), and individual cell counts were obtained (A) as per Ludwig et al. (2006b). No significant differences (P <0.01, t-test) were noted in Oct4 expression or cell proliferation in frozen versus fresh medium. Data represents 3 replicates in triplicate. The medium can be frozen up to 1 year with no decrease in performance. The authors have used frozen media to expand cultures for at least 6 months.

Embryonic and Extraembryonic Stem Cells

1C.2.9 Current Protocols in Stem Cell Biology

Supplement 2

Selenium stock (0.07 mg/ml) Dissolve 7 mg sodium selenite (Sigma) into 100 ml Type I water. Filter sterilize and store the resulting solution in 1-ml aliquots at 4◦ C up to 6 months. CAUTION: Sodium selenite is highly toxic by inhalation. Appropriate safety precautions should be taken when handling, including the use of a respirator. If possible, all work with this compound in its powdered state should be performed in the fume hood.

Stock B Slowly dissolve 67 g of BSA (Sigma; cat. no. A2153) into 500 ml Type I water while gently stirring at room temperature (this step should take up to several hours). To this solution, add:

2.8 g sodium bicarbonate (Sigma) 33 mg thiamine hydrochloride (Sigma) 10 mg reduced glutathione (Sigma) 330 mg L-ascorbic acid 2-phosphate Mg-salt (Sigma) 516 mg γ-aminobutyric acid (GABA, Sigma) 212 mg lithium chloride (LiCl, Sigma) 6.5 µl pipecolic acid stock (see recipe) 1 ml selenium stock (see recipe) 10 ml Trace Mineral Stock B (MediaTech/Cellgro) 5 ml Trace Mineral Stock C (MediaTech/Cellgro) 10 ml human insulin solution (Sigma) 10 ml bFGF stock (see recipe) 10 ml TGFβ stock (see recipe) 10 ml Defined Lipid Concentrate (Invitrogen) Using a Class A volumetric flask (Fisher), bring volume up to 1 liter with Type 1 water. Filter sterilize the resulting solution and store in 200-ml aliquots for up to 2 weeks at 4◦ C, or up to 6 months at −80◦ C. BSA must be added prior to the incorporation of any other protein components. Insulin, in particular, readily adheres to glass, and significant amounts will be lost to the glassware if added prior to BSA. BSA purchased from Sigma has traditionally been used in the authors’ laboratory with good results. Variations have been seen from lot to lot on occasion, however, and therefore batch testing of all serum-derived products is imperative. Be cautious to avoid the formation of bubbles when dissolving, mixing, and filtering this solution. Loss of proteins will result if bubble formation is not adequately controlled. Use only low-protein binding filters, PES membrane, or equivalent.

TGFβ stock (300 ng/ml) Dissolve 30 µg human TGFβ (R&D Systems) into 600 µl 4 mM HCl (Sigma). Add 99.4 ml diluent solution (see recipe). Store 20-ml aliquots up to 6 months at −80◦ C.

Washing medium Add 1.48 g calcium- and magnesium-free DMEM/F-12 (Invitrogen) to 100 ml Type I water and stir until dissolved, adding low heat as necessary. Adjust the pH to 7.2 and the osmolarity to 340 ±5 mOsM using sodium bicarbonate. Filter sterilize the medium, and store in 12- to 60-ml aliquots up to one month at 4◦ C. Warm an aliquot to 37◦ C before use. Defined, FeederIndependent Medium for hESC Culture

1C.2.10 Supplement 2

Current Protocols in Stem Cell Biology

COMMENTARY Background Information Several media have now been reported to support the feeder-independent culture of human ES cells (Table 1C.2.1), two of which have also been reported to support initial derivation (Klimanskaya et al., 2005; Ludwig et al., 2006a). FGF-signaling, and a balance between BMP-family members promoting differentiation and Activin/TGFβ family members inhibiting differentiation are common themes across these different media formulations. TeSR1 is one of the serum-free, animal product-free, defined media that supports both culture and derivation of human ES cells (Ludwig et al., 2006b). Although the initial formulation of TeSR1 contained only recombinant or human-sourced protein components, subsequent modifications include the use of animal sourced proteins (e.g., BSA, Matrigel) and cloned zebrafish bFGF that significantly reduced costs [mTeSR1: (Ludwig et al., 2006a)] and made the medium a practical alternative for most laboratories. The authors have now been using this defined medium for all of their routine human ES cell cultures for more than a year. Human ES cells cultured in mTeSR for >40 passages maintain a normal karyotype, expression of appropriate ES cell markers (>90% Oct4, SSEA4, Tra 1-60, Tra 1-81), and the ability to form all three germ layers in both embryoid bodies and teratomas.

Critical Parameters Traditional serum- and MEF-based human ES cell culture can mask deficiencies within the culture system. In the absence of these components, quality control within the laboratory takes on an increased level of importance. Deficiencies in quality control are rapidly evident in feeder-independent culture systems. Vigilance in monitoring equipment and screening reagents is required to achieve the highest quality and most consistent cultures. Maintenance of sterility is critical. All manipulations must be performed in a Class II Biological Hazard Flow Hood. Furthermore, all hoods should be certified at least yearly to ensure proper flow, and the UV bulbs should be replaced [this will depend on usage, per the manufacturers’ recommendations (total use hours); at a minimum, the UV bulbs should be replaced yearly]. It is difficult to determine if UV lights are emitting the appropriate wavelength to ensure sterilization, and the bulb will continue to illuminate even when the UV is in-

adequate. Replacing the UV bulbs on a regular basis will help to ensure continued sterility of cultures. Previous studies have demonstrated the impact of alterations in the physiochemical environment on human ES cell culture performance (Ludwig et al., 2006b). Minor alterations in pH and osmolarity can dramatically affect the proliferation and differentiation of cell cultures. Changes in CO2 concentrations affect pH, and reductions in environmental humidity can affect osmolarity of media. Cells in culture, specifically undifferentiated and germ cells, can be exquisitely sensitive to temperature variations. Changes in temperature as small as 0.5◦ C can have a dramatic effect on embryo viability in culture (McKiernan and Bavister, 1990; Shi et al., 1998; Abeydeera et al., 2001). While no published studies have investigated the impact of temperature on human ES cell competence, ES cell sensitivity may be similar to that of embryos. Therefore, atmosphere, humidity levels, and temperature of incubators must be monitored daily. Bovine serum albumin and Matrigel are biologically sourced products. While recombinant alternatives to BSA are available, they are beyond the financial reach of the common research laboratory. Studies have demonstrated significant inconsistencies between lots of serum albumin (McKiernan and Bavister, 1992). Individual lots of albumin, therefore, must be carefully screened because they vary considerably in their ability to sustain human ES cell growth. Likewise, in the authors’ laboratory they have noticed variations between lots of Matrigel, and recommend screening each lot to ensure it is sufficient to adequately support undifferentiated human ES cell culture. Water quality is perhaps the most critical factor in the success of any culture system. Only Type I reagent-grade water should be used in the preparation of solutions used for human ES cell culture. While “sterile water” is generally available for purchase, it is not manufactured to meet the appropriate specifications, and is not an adequate substitute for Type I reagent-grade water (Mather et al., 1986). As water quality is significantly affected by storage, even high-quality water should not be stored for an extended period of time (Gabler et al., 1983), but rather used directly from the source. The authors recommend using an inhouse water purification system, such as the Milli-Q Ultrapure Water System (Millipore).

Embryonic and Extraembryonic Stem Cells

1C.2.11 Current Protocols in Stem Cell Biology

Supplement 2

Table 1C.2.1 Feeder-Independent Human ES Cell Culture Systems

Formula Xeno- Medium System disfree defined defined closed

Medium

Basal medium

TLFa

KO-DMEM KOSR

4 ng/ml Fibronectin bFGF, TGFβ

N

N

N

N

Amit et al. (2004)

UMFN

DMEM/F12 KOSR

40 ng/ml bFGF, noggin

Matrigel

N

N

N

N

Xu et al. (2005b)

Ea

KO-DMEM KOSR

40 ng/ml bFGF, Flt3L

Matrigel

N

N

N

N

Xu et al. (2005a)

DSR+ Activin

KO-DMEM KOSR

Activin, KGF

Laminin

N

N

N

N

Beattie et al. (2005)

Unnamedb

KO-DMEM KOSR

10 ng/ml bFGF, 20 ng/ml hLIF

Murine cell extraction

N

N

N

N

Klimanskaya et al. (2005)

SR-bFGF KO-DMEM KOSR

36 ng/ml bFGF

Matrigel

N

N

N

N

Wang et al. (2005)

NC-SFM X-Vivo 10

80 ng/ml bFGF

Laminin

Y

Y

Y

N

Li et al. (2005)

Supplement

—

Key medium Matrix additives

Citation

CDM

IMDM+F12 BSA

12 ng/ml FBS FGF, Activin

N

Y

N

Y

Vallier et al. (2005)

UM100

DMEM/F12 KOSR

100 ng/ml bFGF

N

N

N

N

Levenstein et al. (2005)

TeSR1b

DMEM/F12 HSA

100 ng/ml Matrigel or bFGF, human TGFβ, LiCl, matrix PA, GABA

Y

Y

Y

Y

Ludwig et al. (2006b)

HESCO

X-Vivo

HSA

4 ng/ml bFGF, Wnt3a, April/BAFF

Matrigel or fibronectin

Y

Y

Y

N

Lu et al. (2006)

N2-CDM DMEM/F12

BSA

20 ng/ml bFGF, N2, B27

Matrigel

N

Y

N

N

Yao et al. (2006)

NBF

DMEM/F12

—

100 ng/ml bFGF, N2, B27

Fibronectin

Y

Y

Y

N

Liu et al. (2006)

mTeSR1

DMEM/F12

BSA

100 ng/ml Matrigel bFGF, TGFβ, LiCl, PA, GABA

N

Y

N

Y

Ludwig et al. (2006a)

Matrigel

a ∼20% differentiation. b Supports derivation.

Defined, FeederIndependent Medium for hESC Culture

The system should be maintained regularly, and monitored daily. Total organic content in particular can have a dramatic and devastating effect on culture performance (Fig. 1C.2.4), and may not be detectable at the point of use. Because even carefully maintained systems

can occasionally demonstrate substandard performance, water samples should be sent out regularly for independent testing of sterility, endotoxin, and total organic content. Glassware used for preparation of media should be acid stripped before use, and washed

1C.2.12 Supplement 2

Current Protocols in Stem Cell Biology

Figure 1C.2.4 Effect of water source on human ES cell cultures. Human ES cells were split from the same parental culture into medium made using water source A (A) or water source B (B) and cultured for three passages. All parameters with the exception of water source were identical for both cultures. Both water sources were Milli-Q Ultrapure Water Systems that had been well maintained and were not noticeably different upon observation. At the end of the culture period, cells exposed to water source B demonstrated increased levels of spontaneous differentiation and decreased proliferation. Testing of the water from each source by an outside agency revealed that water from source B contained elevated total organic counts (TOC). Scale bar = 100 µm.

between uses. Soap should never be used to wash glassware, as residues cannot be effectively removed, and will be toxic to human ES cell cultures. Instead, glassware should be rinsed a minimum of 10 times with highquality (Type 1 reagent-grade) water immediately after every use. Glassware should be wrapped in foil and baked dry at a temperature not below 200◦ C. Acid stripping should be repeated every 3 to 5 uses. Glutamine and sodium bicarbonate are highly labile, and are affected by changes in temperature, headspace in containers, and duration of storage. Because of this, media should be kept in an appropriate-sized container to reduce exposure to air, and should not be maintained longer than 2 weeks at 4◦ C. Appropriate passage timing is absolutely critical to the success of this culture system. While other culture methods allow for some flexibility in split timing, in this feederindependent system there is roughly a 24-hour window to passage cells and achieve optimum attachment and continued undifferentiated proliferation. Passage too early and cells will not attach, too late and cultures will easily differentiate. Cultures should be passaged when colony centers become dense, appearing brighter than the edges when viewed using phase contrast microscopy (Fig. 1C.2.2). When initially working with this system, splitting multiple wells of a single culture on successive days and observing the resulting

cultures in the days immediately following passaging may identify appropriate timing. Cells cultured in mTeSR, if maintained properly (i.e., high-quality water and media components, appropriate passage timing, adequate equipment maintenance) should continue to expand with <5% spontaneous differentiation routinely. Autologous feeder formation is not a feature of the mTeSR culture system. Presence of this type of spontaneous differentiation should be viewed as a sign of quality control issues. Cultures that express >10% spontaneous differentiation should not be maintained. “Pick to keep” is not recommended to rescue cultures that have differentiated in any system: co-culture, feeder-free, or feeder-independent. This technique places tremendous selection pressure on cultures and can drive them toward an abnormal karyotype. The only circumstances under which the authors would recommend these kinds of heroic measures are with modified or very low passage cultures that are irreplaceable. Cells cultured in mTeSR medium can be frozen using mTeSR medium supplemented with 20% FBS and 10% DMSO. The authors recommend freezing at twice the standard density. The authors routinely freeze 2 confluent well/vials and thaw 1 vial into 1 well of a 6-well plate. The authors have not tested vitrification. Stem Cell Technologies (the commercial producer of mTeSR) is currently developing an improved freezing medium specifically

Embryonic and Extraembryonic Stem Cells

1C.2.13 Current Protocols in Stem Cell Biology

Supplement 2

Table 1C.2.2 Troubleshooting Guide for Feeder-Independent Culture of Human ES Cells Protocols

Problem

Possible cause

Solution

Low or no attachment at passage

Dispase splitting: Dispase solution may not have been adequately rinsed away.

Increase number and volume of rinses to assure complete removal of dispase solution prior to plating cells.

EDTA splitting: Cells may have been individualized prior to plating.

Decrease time of EDTA incubation and/or decrease disruption of colonies post-incubation.

EDTA splitting: Cell may be damaged by inappropriate osmolarity or pH.

Check and adjust osmolarity and/or pH of solution as appropriate.

Quality of Matrigel may not be appropriate for human ES cell culture.

Screen Matrigel.

Inappropriate passage timing

Passage cells one day later to increase attachment at passage.

Good attachment, Inappropriate pH but limited proliferation

Increased spontaneous differentiation

Check pH of medium post equilibration and adjust accordingly.

Inappropriate osmolarity

Check medium osmolarity and adjust accordingly.

Poor water quality

Test TOC of water (should not exceed 30 ppb).

Small, isolated areas of differentiation, not Screen multiple lots of BSA to assure quality before exceeding 10% of the culture, are normal and use. can be removed by aspiration with a Pasteur pipet at passaging. Increased differentiation may be due to the following causes: Poor quality BSA Poor quality Matrigel

Screen prior to use.

Inadequate water quality

Use only Type 1 reagent-grade water, measuring 18.2 mOhm. Routinely test total organic content and endotoxin levels to assure quality.

Inappropriate passage timing

Passage cells one day earlier to reduce spontaneous differentiation following passage.

for mTeSR cells, and it will be available in October of 2007.

Troubleshooting See Table 1C.2.2 for troubleshooting tips.

Anticipated Results

Defined, FeederIndependent Medium for hESC Culture

Feeder-independent human ES cell culture is relatively simple and efficient provided that the medium is properly prepared. Transfer of cells from MEF-containing cultures to feederindependent conditions may result in some MEF carryover for the first few passages. After the initial passages, however, cultures should remain clear of “feeder-like” cells. Morphologically, cells will appear as expected, exhibiting classic ES cell morphology (minimum of 2 nucleoli, large nucleus to cytoplasm ratio, distinct cell borders; Fig. 1C.2.5). Individual cells, however, will be smaller than in standard

culture conditions, and users may easily underestimate the number of cells within a culture. If cell numbers are important, it is advisable to perform cell counts. Cultures should proliferate well, with <10% spontaneous differentiation overall. On average, 2 to 4 × 106 cells/well can be expected at passage. Generally, cells can be passaged at split ratios between 1:6 and 1:10 every 7 days. The authors have not seen elevated karyotypic instability in this system compared to standard culture systems, and if cells are maintained properly (on-time passaging and limited disruption at passaging), normal karyotypes can be expected. Cells will remain pluripotent, expressing appropriate ES cell markers (Fig. 1C.2.5), and retain the ability to differentiate into all three germ layers. Cloning efficiency is reduced in this culture system however, and users should expect

1C.2.14 Supplement 2

Current Protocols in Stem Cell Biology

Figure 1C.2.5 Morphology and marker expression of human ES cells cultured in mTeSR. Following 15 passages of culture in mTeSR medium on Matrigel, H9 (WA09) human ES cells demonstrate classic ES cell morphology (A), and >95% of cells express Oct4 when stained (B). Scale bar = 10 µm.

<0.1% single cell survival rates using mTeSR medium as formulated. Increasing cloning efficiencies remains an important area of research in human ES cell culture overall, and is particularly important in feeder-independent culture systems.

Time Considerations Media Allow ∼10 to 15 min for the preparation of the following stock solutions: diluent stock, pipecolic acid stock, bFGF stock, TGFβ stock, and L-glutamine stock. When preparing Stock B, allow at least 4 hr for BSA to dissolve. The remainder of the Stock B preparation should take ∼30 min, proceeding slowly to prevent bubble formation. Approximately 30 min should be allowed for assembly of mTeSR culture medium, including pH and osmolarity adjustments.

Passaging Matrigel coating of tissue culture plates will take 3 to 5 min per plate, and must be done at least 1 hour prior to passaging cells. All reagents should be warmed for a minimum of 15 min prior to passaging cells. Allow 1 to 5 min per plate for observation and preparation (marking of differentiation). Once splitting and washing solutions are warmed, allow 7 to 10 min per plate for EDTA passaging, 12 to 15 min per plate for dispase passaging, and 10 to 12 min per plate for collagenase passaging. Culture Medium should be warmed for ∼30 min before feeding cells. Once medium is properly warmed, completing the culture protocol should take no more than 3 to 5 min per plate daily.

Literature Cited Matrigel Thawing bottles of Matrigel will take at least 12 hr and must be performed at 4◦ C on ice. It is most convenient to do this overnight. Allow at least 1 hour for tips and tubes to properly chill before aliquoting Matrigel. Aliquoting Matrigel stock will require ∼1 hr/bottle. While concentrations vary, on average each bottle of Matrigel will cover between 150 and 200 6-well plates.

Abeydeera, L.R., Wang, W.H., Prather, R.S., and Day, B.N. 2001. Effect of incubation temperature on in vitro maturation of porcine oocytes: nuclear maturation, fertilization and developmental competence. Zygote 9:331-337. Amit, M., Shariki, C., Margulets, V., and ItskovitzEldor, J. 2004. Feeder layer- and serum-free culture of human embryonic stem cells. Biol. Reprod. 70:837-845. Beattie, G.M., Lopez, A.D., Bucay, N., Hinton, A., Firpo, M.T., King, C.C., and Hayek, A. 2005.

Embryonic and Extraembryonic Stem Cells

1C.2.15 Current Protocols in Stem Cell Biology

Supplement 2

Activin A maintains pluripotency of human embryonic stem cells in the absence of feeder layers. Stem Cells 23:489-495.

addition of common water contaminants on the growth of cells in serum-free media. BioTechniques 4:56-63.

Buzzard, J.J., Gough, N.M., Crook, J.M., and Colman, A. 2004. Karyotype of human ES cells during extended culture. Nat. Biotechnol. 22:381-382.

McKiernan, S.H. and Bavister, B.D. 1990. Environmental variables influencing in vitro development of hamster 2-cell embryos to the blastocyst stage. Biol. Reprod. 43:404413.

Gabler, R., Hedge, R., and Hughes, D. 1983. Degradation of high purity water on storage. J. Liq. Chromatogr. 6:2565-2570. Klimanskaya, I., Chung, Y., Meisner, L., Johnson, J., West, M.D., and Lanza, R. 2005. Human embryonic stem cells derived without feeder cells. Lancet 365:1636-1641. Levenstein, M.E., Ludwig, T.E., Xu, R.H., Llanas, R.A., Vandenheuvel-Kramer, K., Manning, D., and Thomson, J.A. 2005. Basic FGF support of human embryonic stem cell self-renewal. Stem Cells 24:568-574. Li, Y., Powell, S., Brunette, E., Lebkowski, J., and Mandalam, R. 2005. Expansion of human embryonic stem cells in defined serumfree medium devoid of animal-derived products. Biotechnol. Bioeng. 91:688-698. Liu, Y., Song, Z., Zhao, Y., Qin, H., Cai, J., Zhang, H., Yu, T., Jiang, S., Wang, G., Ding, M., and Deng, H. 2006. A novel chemicaldefined medium with bFGF and N2B27 supplements supports undifferentiated growth in human embryonic stem cells. Biochem. Biophys. Res. Commun. 346:131-139. Lu, J., Hou, R., Booth, C.J.,Yang, S.H., and Snyder, M. 2006. Defined culture conditions of human embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 103:5688-5693. Ludwig, T.E., Bergendahl, V., Levenstein, M.E., Yu, J., Probasco, M.D., and Thomson, J.A. 2006a. Feeder-independent culture of human embryonic stem cells. Nat. Methods 3:637-646. Ludwig, T.E., Levenstein, M.E., Jones, J.M., Berggren, W.T., Mitchen, E.R., Frane, J.L., Crandall, L.J., Daigh, C.A., Conard, K.R., Piekarczyk, M.S., Llanas, R.A., and Thomson, J.A. 2006b. Derivation of human embryonic stem cells in defined conditions. Nat. Biotechnol. 24:185-187. Mather, J., Kaczarowski, F., Gabler, R., and Wilkins, F. 1986. Effects of water purity and

McKiernan, S.H. and Bavister, B.D. 1992. Different lots of bovine serum albumin inhibit or stimulate in vitro development of hamster embryos. In Vitro Cell. Dev. Biol. 28A:154-156. Mitalipova, M.M., Rao, R.R., Hoyer, D.M., Johnson, J.A., Meisner, L.F., Jones, K.L., Dalton, S., and Stice, S.L. 2005. Preserving the genetic integrity of human embryonic stem cells. Nat. Biotechnol. 23:19-20. Shi, D.S., Avery, B., and Greve, T. 1998. Effects of temperature gradients on in vitro maturation of bovine oocytes. Theriogenology 50:667-674. Vallier, L., Alexander, M., and Pedersen, R.A. 2005. Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell. Sci. 118:44954509. Wang, L., Li, L., Menendez, P., Cerdan, C., and Bhatia, M. 2005. Human embryonic stem cells maintained in the absence of mouse embryonic fibroblasts or conditioned media are capable of hematopoietic development. Blood 105:45984603. Xu, C., Rosler, E., Jiang, J., Lebkowski, J.S., Gold, J.D., O’Sullivan, C., Delavan-Boorsma, K., Mok, M., Bronstein, A., and Carpenter, M.K. 2005a. Basic fibroblast growth factor supports undifferentiated human embryonic stem cell growth without conditioned medium. Stem Cells 23:315-323. Xu, R.H., Peck, R.M., Li, D.S., Feng, X., Ludwig, T., and Thomson, J.A. 2005b. Basic FGF and suppression of BMP signaling sustain undifferentiated proliferation of human ES cells. Nat. Methods 2:185-190. Yao, S., Chen, S., Clark, J., Hao, E., Beattie, G.M., Hayek, A., and Ding, S. 2006. Long-term selfrenewal and directed differentiation of human embryonic stem cells in chemically defined conditions. Proc. Natl. Acad. Sci. U.S.A. 103:69076912.

Defined, FeederIndependent Medium for hESC Culture

1C.2.16 Supplement 2

Current Protocols in Stem Cell Biology

Isolation and Propagation of Mouse Embryonic Fibroblasts and Preparation of Mouse Embryonic Feeder Layer Cells

UNIT 1C.3

Anna E. Michalska1 1

Monash University, Victoria, Australia

ABSTRACT To realize their potentials, embryonic stem (ES) cells must be maintained in optimal culture conditions that preserve their pluripotency and self-renewal capacity. Mouse embryonic fibroblasts (MEFs) are used to prepare a feeder cell layer that supports the growth of ES cells and the quality of feeders is crucial for the maintenance of undifferentiated ES cells in prolonged culture. The protocols provided in this unit describe aspects of isolation and expansion of MEFs and maintenance of established feeder cells. Preparation of mitotically inactivated feeder cell layer (treatment with mitomycin C or γ-irradiation) is also described. In addition, a method for counting cell numbers and a simple method for detection of mycoplasma contamination by in situ DNA staining are also provided. Methodology described has been tested in a real laboratory environment and provides detailed information regarding resource and time requirements as well as critical parameters and troubleshooting. Curr. Protoc. Stem Cell Biol. 3:1C.3.1-1C.3.17. C 2007 by John Wiley & Sons, Inc.
Keywords: embryonic stem (ES) cells r culture r mouse embryonic fibroblasts (MEFs) r isolation and expansion r feeder cell layer r MEFs freezing and thawing r mycoplasma detection r protocols and methods

INTRODUCTION Mouse embryonic fibroblasts (MEFs) are primary cell lines derived from mouse fetuses between 12.5 and 14.5 days of gestation. These cells are easy to isolate and propagate in culture and can be expanded to large numbers. MEFs are routinely used to produce feeder cell layers that support the growth of a variety of fastidious cultured cell types, including stem cells. Feeder cells are produced by inhibition of cell division with mitomycin C (MMC) or γ-irradiation. Following the treatment, which inhibits their division, the feeder cells remain metabolically active. The function of feeders is to provide growth support for the cells plated on them. Both MEFs and feeder cells can be frozen and stored indefinitely in liquid nitrogen. When thawed, they remain viable and can be further expanded (MEFs) or used for the preparation of feeder layers. The protocols provided in this unit describe aspects of isolation and expansion of MEFs (Basic Protocol 1), and freezing and thawing of MEFs and inactivated feeder cells (Support Protocols 1 and 2). In addition, a simple method for mycoplasma detection by in situ DNA staining is described (Support Protocol 3). Preparation of feeder cell layer by treatment with mitomycin C (MMC) is described in Basic Protocol 2, while treatment by γ-irradiation is described in Alternate Protocol. A method for counting cell number is also provided (Support Protocol 4). NOTE: It is expected that the researcher has access to tissue culture laboratory and equipment such as: class II biological hazard hood or laminar flow horizontal draft Current Protocols in Stem Cell Biology 1C.3.1-1C.3.17 Published online October 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c03s3 C 2007 John Wiley & Sons, Inc. Copyright


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hood, humidified 37◦ C 5% CO2 incubator, low-speed centrifuge, water bath, liquid nitrogen storage facility, −80◦ C freezer, inverted phase contrast microscope, fluorescence microscope, and other standard laboratory equipment. NOTE: All techniques involving media/reagents preparation and cell manipulation must be performed under sterile conditions unless specified otherwise. NOTE: All manipulations and waste disposal must be carried out according to the institutional biohazard guidelines. NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Committee for Ethical Animal Care and Use (IACUC) and must conform to government regulations for the care and use of laboratory animals. BASIC PROTOCOL 1

ISOLATION AND PROPAGATION OF MOUSE EMBRYONIC FIBROBLASTS Mouse embryonic fibroblasts (MEFs) are used to prepare a feeder cell layer that supports the growth of embryonic stem (ES) cells. MEFs are isolated from 12.5 to 14.5 day old mouse fetuses (the morning the copulation plug is found is designated as day 0.5). Time-mated female mice can be purchased from a commercial source or can be prepared in-house. It is recommended that mice are mated naturally rather than following superovulation (superovulation can result in a large number of very small fetuses that will produce inferior feeder cells). Once expanded, MEFs can be frozen and stored for later use. All newly isolated MEF cultures should be tested for the presence of mycoplasma.

Materials 12.5 to 14.5 days post-coitum (dpc) pregnant female mouse 70% (v/v) ethanol (see recipe) Phosphate-buffered saline, calcium and magnesium free (CMF-PBS; see recipe), ice cold and room temperature 0.25% (w/v) trypsin/EDTA solution for fetal tissue digestion (see recipe) MEF culture medium (see recipe) 0.05% (w/v) trypsin/EDTA solution for MEFs passaging (see recipe) Pair of scissors and forceps (do not have to be sterile) Sterile instruments (iris scissors, 2 pairs of watchmaker forceps no. 5, scalpel blade) 10-cm bacteriological petri dishes 5- and 10-ml plastic pipets 50-ml centrifuge tubes 37◦ C water bath 37◦ C, 5% CO2 incubator 75-cm2 tissue culture flasks Inverted microscope Additional reagents and equipment for euthanasia by cervical dislocation (Donovan and Brown, 2006) Collect fetuses 1. Kill pregnant female mouse by cervical dislocation (Donovan and Brown, 2006) or other IACUC method. Steps 1 to 7 can be performed on the bench. CMF-PBS should be ice cold to anesthetize dissected fetuses. Isolation and Preparation of MEFs

2. Submerge the whole mouse for a few seconds in 70% ethanol to sterilize. Lay the mouse on its back.

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3. Lift the abdominal skin with forceps and make a small transverse incision with a pair of scissors (forceps and scissors do not have to be sterile). Grasp the skin above and below the cut with your fingers and pull apart to expose the muscles of the entire abdomen. 4. With the set of sterile instruments cut open the abdominal wall and lift intestines to expose uterus. 5. Using forceps, grasp the vagina end of the uterus, cut the uterine horns below uterovaginal junction and dissect from mesometrium and cut off any fat tissue attached. 6. Cut each uterine horn below the ovary and place in a 10-cm bacteriological petri dish containing 10 ml of ice-cold CMF-PBS. Swirl around to remove blood. 7. Transfer the uterus to the second petri dish containing ice-cold CMF-PBS and at this stage move the dish into a laminar flow hood. 8. Cut open the uterine wall with fine scissors and release the embryos together with placenta and surrounding membranes into CMF-PBS. Discard the uterus. Collect only healthy looking fetuses and discard those that appear degenerated or undergoing resorption.

Dissect fetuses 9. With fine watchmakers forceps, separate each embryo from its placenta and membranes, decapitate (optional) by pinching off the head with forceps and remove visceral tissues (mostly liver and heart which appear red). This can be done by securing the carcass on its back with one pair of forceps and scooping the viscera with the second pair. Although not necessary, this procedure can be easier to perform (especially for a novice) under the dissecting microscope. It is not crucial to remove all of visceral tissue. These cells will not survive subsequent subculture.

10. Transfer the fetal tissue into a clean dish. 11. Wash the fetal tissue in three further changes of ice-cold CMF-PBS to remove as much blood as possible.

Prepare MEFs 12. Mince the tissue using iris scissors or a scalpel blade in a minimal volume of CMFPBS. 13. Transfer minced tissue into 50-ml tube with a 5-ml pipet. 14. Add 0.5 to 1.0 ml of 0.25% trypsin/EDTA solution per embryo and incubate 10 to 15 min in the water bath at 37◦ C with gentle agitation. 15. Pipet the tissue slurry with a 5-ml pipet until few chunks remains. DNase I (0.1 µg/ml) can be added to eliminate clumping due to the release of highmolecular-weight DNA from dead cells.

16. Add ∼2 vol of MEF culture medium to neutralize trypsin and pipet again. Following trypsin treatment, the tissue suspension should be free of any large pieces. It should not be too viscous—high viscosity is due to extensive cell damage and release of DNA. If a lot of large pieces are still present, they can be removed by allowing them to settle to the bottom of the tube for 1 to 2 min. Embryonic and Extraembryonic Stem Cells

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17. Transfer aliquots equivalent to 1.5 to 2 fetuses into 75-cm2 tissue culture flasks containing 20 to 30 ml of MEF culture medium. This is passage 0 (P0). There is no need to centrifuge the cells to remove trypsin/EDTA. Serum supplementing MEF culture medium contains sufficient trypsin inhibitor and EDTA is diluted in a large volume of culture medium. As a guide, 2, 1.5, and 1 fetuses from 12.5 dpc, 13.5 dpc, and 14.5 dpc female, respectively are plated per 75-cm2 culture flask.

18. Incubate the cells at 37◦ C in 5% CO2 incubator. 19. The following day change the medium to remove cell debris and unattached tissue. At this stage many cell types might be seen in addition to fibroblasts (e.g., blood, nerve, cartilage). These other cells will not survive the subsequent subculture.

20. When the cultures become 80% to 90% confluent (this should be in a day or two), expand cells (see below) or freeze them (see Support Protocol 1).

Propagate MEFs 21. Aspirate and discard MEF culture medium. Wash the cell surface with 10 ml CMFPBS at room temperature. 22. Add 1.5 ml of 0.05% trypsin/EDTA and swirl to ensure the entire cell surface is covered with the solution. 23. Incubate 3 to 5 min at room temperature. 24. Slightly tap the flask to dislodge cells; avoid splashing cells onto sides of flask. Check under inverted microscope to ensure that all cells have detached. 25. Add 8.5 ml of fresh MEF culture medium to neutralize trypsin. Pipet the cell suspension vigorously with a 10-ml pipet to wash the cells off the bottom of the flask and to produce a single-cell suspension. 26. Divide the cell suspension into four to five fresh 75-cm2 tissue culture flasks (split ratio 1:4 to 1:5) containing 20 to 30 ml MEF culture medium. This is passage 1 (P1). 27. Incubate the cells in 37◦ C, 5% CO2 incubator until cultures become 80% to 90% confluent (2 to 3 days) and passage following steps 21 to 26. The volumes given are for a 75-cm2 tissue culture flask and should be adjusted accordingly for smaller or larger flasks. To ensure a sufficient number of cells for long-term usage, MEFs can be expanded for three to five passages before being used for feeder layer preparation. Stocks of MEFs can be frozen at each passage and stored for later use (see Support Protocol 1). The split ratio will depend on a number of factors such as culture confluency, cell morphology, and passage number. MEFs should not be plated too sparsely or allowed to become overgrown. MEFs will grow slower with each passage and became senescent after 20 cell divisions (∼5 to 7 passages). MEFs at an early passage (P0 to P2), 80% to 90% confluent and with epithelial, pavement-like appearance can be split at 1:4 to 1:6 ratio. MEFs at later passages can be split at 1:3 or 1:2 ratio. Better indication of plating density, but less convenient, is to count the cells and plate at 1.7 to 2.6 × 104 cells per cm2 . SUPPORT PROTOCOL 1

Isolation and Preparation of MEFs

FREEZING MEFS Expanded early passage (P2, P3) MEFs can be frozen and stored in liquid nitrogen for future use. Devices that allow controlled freezing can be used if available. The simple method described below provides 1◦ C/min cooling rate from room temperature to −80◦ C and gives reliable results.

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Materials 75-cm2 flask(s) of 80% to 90% confluent MEFs (Basic Protocol 1) Freezing solution (see recipe) Liquid nitrogen 1-ml cryovials 15-ml centrifuge tubes Container with ice Freezing container (e.g., Mr. Frosty or Styrofoam box; Nalgene) Additional reagents and equipment for removing cells from the flasks (Basic Protocol 1) 1. Label cryovials with the name of the cell line, passage number, and number of cells or surface area. Keep freshly prepared freezing solution on ice. 2. Follow steps 21 through to 25 of Basic Protocol 1 to remove the cells from the flask. 3. Transfer the cell suspension into a 15-ml centrifuge tube. At this stage take an aliquot to count the cell number (see Support Protocol 4). 4. Centrifuge the cell suspension 5 min at 270 × g, room temperature. 5. Carefully aspirate supernatant without touching the cell pellet. Tap the tube gently to disperse the pellet. 6. Resuspend the cell pellet in a cold freezing solution at the concentration of 2 to 10 × 106 /ml and transfer 1-ml aliquots into each cryovial. Freezing solution contains DMSO that is toxic to cells at room temperature. Keep the cells on ice before transferring to freezing container. The concentration of frozen MEFs will depend on the number of harvested cells and the requirements of a particular laboratory. It is convenient to freeze between 2 to 10 × 106 cells per cryovial since upon thawing ∼2 × 106 cells are plated into each 75-cm2 tissue culture flask.

7. Place cryovials at −80◦ C freezer in a Mr. Frosty or Styrofoam box. 8. The following day transfer the vials into liquid nitrogen for long-term storage.

THAWING MEFS Stocks of active or inactivated MEFs can be stored indefinitely in liquid nitrogen. Following thawing, MEFs can be further expanded and inactivated fibroblasts can be used for feeder layer preparation.

SUPPORT PROTOCOL 2

Materials MEF culture medium (see recipe) Cryovials containing frozen MEFs (Support Protocol 1) 70% (v/v) ethanol (see recipe) 15-ml centrifuge tubes 37◦ C water bath 75-cm2 tissue culture flasks 1. Place 9 ml of MEF culture medium (warmed to 37◦ C) into 15-ml centrifuge tube. Embryonic and Extraembryonic Stem Cells

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2. Remove the frozen cryovial of MEFs from liquid nitrogen storage and place into a 37◦ C water bath. Safety glasses or a face shield mask must be worn when removing vials from liquid nitrogen storage and when thawing cells. When removing from the liquid nitrogen storage, place the vial on dry ice first to allow for any liquid nitrogen trapped inside the vial to evaporate.

3. Thaw as quickly as possible and remove from the water when a small piece of ice remains (∼1 to 2 min). Sterilize the outside of the vial with 70% ethanol. 4. Transfer the contents of the vial dropwise into the 15-ml centrifuge tube containing MEF culture medium. 5. Centrifuge the cell suspension at 270 × g for 5 min, room temperature. 6. Carefully aspirate supernatant without touching the cell pellet. Tap the tube gently to disperse the pellet. Resuspend the cells in 5 ml MEF culture medium and count the number of viable cells (see Support Protocol 4). Plate ∼2 × 106 cells into a 75-cm2 tissue culture flask. 7. Incubate the cells in a 37◦ C,5% CO2 incubator until cultures become 80% to 90% confluent. 8. Passage following steps 21 to 27 of Basic Protocol 1 or use for feeder preparation (see Basic Protocol 2). Following freezing and thawing some cell death occurs. Therefore it is recommended that the cells are counted before plating. In general, recovery of viable cells after freezing should be >80%. SUPPORT PROTOCOL 3

MYCOPLASMA TESTING Prior to use, each isolated batch of MEFs should be routinely tested for the presence of mycoplasma. Mycoplasmas are small (0.2 to 2.0 µm in diameter), self-replicating organisms that in cell culture attach to the cell membrane. Mycoplasma contamination can have detrimental effect on cultured cells since the infection alters all cellular processes leading to diminished cell growth and eventually cell death. There are a number of methods to test for mycoplasma infection, including PCR, biochemical assays, ELISA, or immunofluorescence with a number of detection kits available commercially. Described below is a simple procedure for mycoplasma detection by in situ DNA fluorescence.

Materials Cultures of MEFs MEF culture medium (see recipe) with and without antibiotics Fixative solution (see recipe) Phosphate-buffered saline, calcium and magnesium free (CMF-PBS; see recipe) Hoechst 33258 dye solution (see recipe) 6-cm tissue culture petri dishes Cover slips UV fluoresence microscope Prepare MEFs 1. Grow MEFs for two passages (∼1 week) in MEF culture medium without antibiotics. Follow steps 21 to 27 of Basic Protocol 1. Isolation and Preparation of MEFs

Mycoplasma contamination cannot be prevented or eliminated by the addition of antibiotics, such as penicillin and streptomycin, to the culture medium. However, the presence of antibiotics can mask bacterial contamination.

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2. Plate MEFs at the regular passage density (∼3 × 105 ) onto a sterile coverslip placed in a 6-cm tissue culture petri dish containing 5 ml of MEF culture medium and incubate until cells reach 30% to 50% confluence.

Fix cells 3. Prepare the fresh fixative solution on the day of use. Fixative solution contains glacial acetic acid and steps 3 to 6 should be performed in a chemical fume hood.

4. Without removing culture medium, add ∼5 ml of fixative solution to a dish and leave for 2 min. 5. Decant fixing solution from a dish and add 5 ml of fresh fixative. Leave for 5 min and repeat once more. 6. Decant fixative and air dry growth surface completely if the plate is to be stored (samples may be accumulated at this stage and stained later). 7. If proceeding directly, wash off fixative with 5 ml of CMF-PBS.

Add dye to cells 8. Add 5 ml of Hoechst 33258 dye solution and leave for 10 to 20 min at room temperature in the dark. 9. Discard the Hoechst 33258 dye solution and rinse the plate twice with CMF-PBS. 10. Mount a cover slip with drop of CMF-PBS. 11. Examine the cells under UV fluorescence at 200× to 400× magnification. The presence of mycoplasma is manifested by flecks of fluorescent material attached to the cell membrane and/or in cytoplasm. In uncontaminated cultures, only cell nuclei are stained.

12. Discard infected cultures immediately.

PREPARATION OF MOUSE EMBRYONIC FEEDER CELL LAYERS BY MITOMYCIN C TREATMENT

BASIC PROTOCOL 2

In order to prepare feeder cell layers, MEFs must be mitotically inactivated. This prevents division of the fibroblasts while ensuring that they remain metabolically active. Mitotic inactivation is typically achieved by treatment with mitomycin C (MMC) or by exposure to γ-irradiation (see Alternate Protocol). Mitomycin C treatment is convenient and does not require sophisticated equipment (γ-irradiator); however, MMC is toxic to cultured cells and humans. Care must be taken when preparing MMC solution and the fibroblasts must be extensively washed to prevent toxic effect to ES cells. MEFs between passage 2 and 5 can be used for inactivation and feeder preparation.

Materials 75-cm2 flask(s) of 80% to 90% confluent MEFs (Basic Protocol 1) Mitomycin C (MMC) solution (see recipe) MEF culture medium (see recipe) 0.1% (w/v) gelatin solution (see recipe) Phosphate-buffered saline, calcium and magnesium free (CMF-PBS; see recipe) 0.05% (w/v) trypsin/EDTA solution (see recipe) 37◦ C, 5% CO2 incubator Appropriate tissue culture plates, dishes, or flasks for growing ES cells Inverted microscope

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10-ml plastic pipets 15-ml centrifuge tubes 37◦ C, 5% CO2 incubator Treat MEFs with mitomycin C 1. Aspirate the culture medium from the flask of MEFs and replace with 10 ml fresh MEF culture medium containing 10 µg/ml mitomycin C. 2. Place the flask in a 37◦ C, 5% CO2 incubator and incubate between 2 to 3 hr (minimum for 2 hr). 3. In the meantime coat appropriate tissue culture plates, dishes, or flasks in which ES cells are to be grown with 0.1% gelatin solution. Add enough of the gelatin solution to cover the growth area, leave at room temperature for ∼1 hr and aspirate.

Wash MEFs 4. Aspirate the mitomycin C medium from the MEFs. Wash the cells once with 10 ml of MEF culture medium and twice with 10 ml CMF-PBS. Medium containing mitomycin C has to be disposed as hazardous waste according to institutional health and safety guidelines.

Trypsinize the cells 5. Add 1.5 ml of 0.05% trypsin/EDTA and swirl to ensure the entire cell surface is covered with the solution. 6. Incubate 5 min at room temperature. 7. Slightly tap the flask to dislodge cells (avoid splashing cells onto sides of flask). Check under inverted microscope to ensure that all cells have detached. 8. Add 8.5 ml of MEF culture medium to neutralize trypsin. Vigorously pipet the cell suspension with 10-ml pipet to wash the cells off the flask and to produce a single-cell suspension. 9. Transfer the cell suspension into a 15-ml centrifuge tube. 10. Centrifuge the cell suspension 5 min at 270 × g, room temperature.

Plate feeder cells 11. Carefully aspirate the supernatant without touching the cell pellet. Tap the tube gently to disperse the pellet. Resuspend the pellet in 10 ml of MEF culture medium and count the cell number (see Support Protocol 4). 12. Plate MEFs onto gelatin-coated plates, dishes, or flasks at appropriate cell number and in appropriate volume of MEF culture medium. As a guide, feeder cells should be plated at the following density: 1.0 to 1.5 × 105 /cm2 for mouse ES cells; 2 or 6×104 cells/cm2 for human ES cells (dependent on culture conditions). When receiving ES cells, supplier’s recommendations should always be followed.

13. Place the plates in a 37◦ C, 5% CO2 incubator. MEFs will attach within 20 min and spread out within 12 hr.

14. Prior to plating ES cells, check the plates and change medium to appropriate ES culture medium. Isolation and Preparation of MEFs

The feeder layers can be used the following day and up to 1 week.

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15. Alternatively, freeze the inactivated MEFs following Support Protocol 1, steps 4 through 8. Inactivated cells should be frozen at a slightly higher density to allow for some cell loss (∼20%) following the freezing/thawing procedure.

PREPARATION OF MOUSE EMBRYONIC FEEDER CELL LAYERS BY γ -IRRADIATION

ALTERNATE PROTOCOL

MEFs can be inactivated by exposure to γ-irradiation. This procedure is as effective as MMC treatment and might be a preferred method of inactivation if access to irradiation source is available. The advantage of γ-irradiation is that no toxic reagent is used. However, dose of required irradiation may vary for different cell lines and should be determined by performing a radiation titration.

Materials 75-cm2 flask(s) of 80% to 90% confluent MEFs (Basic Protocol 1) 0.1% (w/v) gelatin solution (see recipe) MEF culture medium (see recipe) Appropriate tissue culture plates/dishes/flasks for growing ES cells 50-ml centrifuge tubes Controlled cesium source for γ-irradiation Additional reagents and equipment for detaching MEFs (Basic Protocol 1) 1. Coat appropriate tissue culture plates, dishes or flasks with gelatin as described in Basic Protocol 2, step 3. 2. Detach MEFs following steps 21 through to 25 of Basic Protocol 1. 3. Transfer the cell suspension into 50-ml centrifuge tube. MEFs from multiple flasks can be pooled into 50-ml tube for γ-irradiation.

4. Centrifuge the cell suspension 5 min at 270 × g, room temperature. 5. Carefully aspirate supernatant without touching the cell pellet. Tap the tube gently to disperse the pellet. Resuspend the cell pellet in 25 ml of MEF culture medium and count the cell number (see Support Protocol 4). 6. Expose MEFs to 30 to 100 Gy (3,000 to 10,000 rads) from a controlled γ-irradiation source. 7. Plate MEFs as described in Basic Protocol 2, steps 11 and 12. 8. Alternatively freeze irradiated MEFs following Support Protocol 1, steps 4 through to 8.

COUNTING NUMBER OF CELLS The growth rate of MEFs depends on the initial plating density. If the cells are seeded at low density, they will grow very slowly, while cells seeded at high density grow at a faster rate and must be passaged more often. In either case the lifespan of MEFs will be reduced. Plating density is also important for feeder layer preparation and will profoundly affect the growth of ES cells. If the feeder cells are too sparse, ES cells will differentiate, and if too dense, feeder layer may detach after a few days resulting in the loss of ES cell culture. The cell number can be counted using an electronic particle counter (e.g., Coulter Counter) or, as described below, using a hemacytometer. In addition, the cell viability can be determined by adding trypan blue solution to cell sample.

SUPPORT PROTOCOL 4

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Materials 0.1- to 0.2-ml-size aliquot of MEFs 0.4% (w/v) trypan blue solution (see recipe) Hemacytometer and coverslip Piston-driven air displacement pipet (e.g., Gilson pipet) 1. Place a coverslip onto counting area (grid) of hemacytometer. 2. Mix the MEF suspension gently and add an equal volume of trypan blue solution (e.g., 0.1 to 0.2 ml of each). 3. Mix the cell sample thoroughly and carefully introduce it underneath the coverslip with a pipet. This is best done by putting the pipet tip at the junction between the hemacytometer and a coverslip and releasing a small volume (10 to 15 µl) of cell suspension—the fluid will be drawn into the chamber by capillary action.

4. Place the hemacytometer under the microscope and focus on the counting grid under 40× total magnification. The grid is divided into 9 large squares.

5. Count the number of cells (Phelan, 2006) in the four corner squares which are divided into 16 smaller squares each. Count the cells lying on the right-hand side and on the bottom lines of the grid in each square and exclude those lying on the left-hand side and top lines. 6. Count clear, refractive cells and larger, dark blue cells separately. The viable cells actively expel the stain and remain round and refractive while dead cells become larger and are stained dark blue.

7. Calculate number of viable cells using the following formula:

N/4 × D × 104 = viable cells/ml, where N is the number of live (unstained) cells counted in four squares, and D is the dilution factor in trypan blue. Multiply the number of cells/ml by the volume of cell suspension to obtain the total number of viable cells. 8. Calculate cell viability as follows: number of live cells/number of live and dead cells × 100% = % viability.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Ethanol, 70% (v/v) 700 ml ethanol, 96%, technical grade 300 ml distilled water Store up to 6 months at room temperature Fixative solution Isolation and Preparation of MEFs

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3 parts absolute ethanol (12 ml) 1 part glacial acidic acid (4 ml) Prepare fixative solution fresh on the day of use CAUTION: Prepare in a chemical fume hood. Current Protocols in Stem Cell Biology

Freezing solution 90% (v/v) fetal bovine serum (FBS) 10% (v/v) dimethyl sulfoxide (DMSO) Prepare freezing solution just prior to use and store on ice Gelatin, 0.1% (w/v) 0.4 g of gelatin powder (e.g., Sigma) 400 ml distilled water Sterilize by autoclaving Store up to 6 months at room temperature Hoechst 33258 dye solution 100 mg Hoechst powder (e.g., Invitrogen, Sigma) 10 ml distilled water Store at 4◦ C protected from light for up to 6 months. For long-term storage store aliquots at −20◦ C. Use at the final concentration of 0.2 µg/ml. CAUTION: Hoechst 33258 dye is a known mutagen. Refer to product data sheet from the manufacturer for handling instructions.

MEF culture medium Dulbecco’s modified Eagle medium (DMEM) with high glucose (4500 mg/liter; e.g., Invitrogen) 10% (v/v) FBS 2 mM L-glutamine or glutaMAX-1 (from 100× stock solution; e.g., Invitrogen) 0.1 mM 2-mercaptoethanol (e.g., Invitrogen) 50 U/mM penicillin, 50 mg/ml streptomycin (from 100× stock solution; Invitrogen) Store up to 2 weeks at 4◦ C After passage 1, use MEF culture medium without antibiotics for propagation.

Mitomycin C solution 2 mg ampule of mitomycin C powder (Sigma) 4 ml sterile distilled water Using a 5 ml syringe and an 18-G needle inject water into an ampule (insert a second needle to vent the ampule). Mix to dissolve and draw back the solution into syringe. Sterilize by filtration through 0.22-µm pore filter. Store solution up to 2 weeks at 4◦ C protected from light. Use at the final concentration of 10 µg/ml. CAUTION: Mitomycin C is a toxic substance. Refer to product data sheet from the manufacturer for handling instructions.

Phosphate-buffered saline calcium and magnesium free (CMF-PBS) PBS tablets or powder (e.g., Sigma, Invitrogen, Calbiochem) Distilled water Sterilize by autoclaving. Store up to 6 months at room temperature Commercially prepared CMF-PBS solution is available (e.g., Invitrogen, Sigma).

Trypan blue, 0.4% (w/v) 0.4 g trypan blue powder (e.g., BDH Laboratory Supplies, Sigma) 0.9 g NaCl 100 ml distilled water

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Pass through 0.45-µm pore filter. Sterilize by filtration through 0.2-µm pore filter Store up to 1 year at room temperature Commercially prepared 0.4% trypan blue solution is available (e.g., Sigma or Invitrogen).

Trypsin/EDTA, 0.25% (w/v) 0.25% (w/v) trypsin/1 mM EDTA solution is purchased from a commercial supplier (e.g., Invitrogen) Store in aliquots at 4◦ C for up to 1 week or at −20◦ C for up to 6 months Avoid repeated freezing and thawing.

Trypsin/EDTA, 0.05% (v/v) 10 ml of 0.25% (w/v) trypsin/EDTA 40 ml CMF-PBS Store in 10-ml aliquots at 4◦ C for up to 1 week or at −20◦ C for up to 6 months Avoid repeated freezing and thawing.

COMMENTARY Background Information

Isolation and Preparation of MEFs

Mammalian embryonic stem (ES) cells, isolated from a morula- or blastocyst-stage embryo, can be grown in vitro indefinitely. The capacity of the ES cells to differentiate in vitro and in vivo into cells from all three germ layers makes them invaluable to study cell differentiation and development, provides models of embryogenesis, and generates cells suitable for drug development and toxicology testing. To realize these potentials, ES cells must be maintained in optimal culture conditions that preserve their pluripotency and self-renewal capacity. The concept of “feeder layers” was developed over 50 years ago when Puck and Marcus (1955) had shown that mitotically inactive fibroblast cells support the clonal derivation of HeLa cells. Such feeders were later used to support clonal propagation of pluripotent mouse embryonal carcinoma (EC) cells. Most EC cells grown in specified culture conditions without feeders have limited differentiation potential, but when grown on feeders, multiple clonal lines with extensive differentiation potentials can be established (Martin and Evans, 1975). When cells were removed from feeders they differentiated into a wide variety of cell types. This provided the concept that feeder cells prevent cells from differentiation and allow them to multiply in vitro in an undifferentiated state. Derivation and maintenance of mouse ES (mES) cells evolved form earlier work on EC cells and the first mES cells were isolated on such feeders (Evans and Kaufman, 1981; Martin, 1981). Extrapola-

tion of this work led to isolation, nearly two decades after, of human ES cells (Thomson et al., 1998; Reubinoff et al., 2000). Embryonic stem cells from a number of species other than mouse and human have been derived using feeder cell support (Gardner, 2004 and references within). The first mES cells were isolated on inactivated, permanent STO fibroblast cell line (derived from SIM strain of mice and thioguanineand ouabain-resistent; Ware and Axelrad, 1972). Other established embryonic fibroblast cell lines, such as C3H 10T1/2 and BALB3T3/A31 were also shown to have the capacity to maintain undifferentiated mES or EC cells in culture (Ogiso et al., 1982; Rathjen et al., 1990). However, a number of researchers preferred to use primary mouse embryonic fibroblasts (MEFs) isolated from fetuses between 12.5 and 18 days of gestation. It has been claimed that primary cells are superior to permanent STO cells and that culture on STO is not successful, or results in karyotypic abnormalities and reduced differentiation ability of mES (Wobus et al., 1984; Doetschman et al., 1985; Suemori and Nakatsuji, 1987). The inconsistency between reports on suitability of STO might have resulted from changes in properties of these cells during culture or from variations in their treatment in different laboratories. An advantage of using MEFs is that they are easy to isolate and, as a primary cell line, seem to produce a more potent, reliable, and reproducible source of feeders. However, the major disadvantage is their limited lifespan as they can undergo only ∼20

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Current Protocols in Stem Cell Biology

divisions before entering senescence (Karatza and Shall, 1984). It has to be pointed out that in standard culture conditions, fibroblasts are grown under oxidative conditions in oxygen concentration of 21%, which is several times greater than that occurring in vivo. It has been shown that the lifespan of primary fibroblasts can be increased (more population doublings) when the oxygen concentration is reduced to a more physiological level of 3% (Parrinello et al., 2003). Since there is also an increased body of evidence on the effects of physiological oxygen levels on embryonic stem cells (Ezashi et al., 2005; Forsyth et al., 2006; Wang et al., 2006), culture in reduced oxygen tension should be considered for the isolation, expansion, and propagation of both types of cells. Although proliferating feeder cells have been used to support the growth and maintenance of ES cells (Xie et al., 2005), a more common approach is to use mitotically inactive feeders. In order to perform their function, the feeder cells must remain metabolically active. The mitotic inactivation is usually achieved by γ-irradiation or mitomycin C (MMC) treatment. Gamma-irradiation causes breaks in DNA double strands while MMC, a chemotherapeutic agent, has the ability to cross-link DNA with high efficiency and to prevent DNA double strand separation during replication. In either case, the synthesis of RNA and proteins continues and feeder cells maintain expression of specific ligands, cytokines, and growth factors. Both methods seem to be equivalent but it has been reported that MMC treatment can alter metabolic state or reduce the metabolic rate of feeder cells (Roy et al., 2001). On the other hand, others have demonstrated that irradiation can decrease the feeder cells lifespan as they tend to detach after extended time in culture (Ponchio et al., 2000). In the end, the method of choice is usually dictated by the convenience and availability of specialized irradiation equipment. Fibroblast feeder cells produce extracellular matrices and a number of soluble factors that are important for proliferation and maintenance of undifferentiated ES cells. The main factor secreted by mouse feeder cells that is responsible for supporting mES cells is cytokine leukemia inhibitory factor (LIF) and mES can be grown in the absence of feeders in a medium containing serum and supplemented with LIF (Smith et al., 1988; Williams et al., 1988). LIF acts by activating JAK/Stat3 signaling pathway and activated Stat3 seems to be sufficient for the maintenance of undiffer-

entiated mES cells (Niva et al., 1998; Matsuda et al., 1999). However, LIF alone can not support clonal growth of mES cells in feeder- and serum-free conditions (Ogawa et al., 2004). It has been postulated that other factors, provided by feeder cells and/or added serum, such as bone morphogenic protein (BMP) and Wnt act synergistically with LIF to maintain mES cell self-renewal (Ying et al., 2003; Ogawa et al., 2006). It is fortuitous that hES cells can be derived and maintained on mouse feeder cells since the action of MEFs in supporting mouse and human ES cells differs. Unlike mouse ES cells, human ES cells seem not to require LIF for their propagation and maintenance of pluripotentiality while BMP2 or BMP4 induces hES differentiation to primitive endoderm or trophectoderm, respectively (Xu et al., 2002; Daheron et al., 2004; Pera et al., 2004). Subsequently it has been found that a number of human-derived feeder cells can support isolation and maintenance of hES cells (see Mallon et al., 2006 for references). Despite many years of using mouse feeder cells, it is still unclear what essential factors they produce that sustain hES cell pluripotency in culture. Preliminary proteomic analysis of MEF-conditioned medium identified 136 proteins that may play a role in maintaining hES cells (Lim and Bodnar, 2002). These include intracellular proteins, extracellular matrix, as well as surface membrane proteins. Comparative proteomic analysis of proteins expressed by MEFs before and after irradiation identified a number of protein species that are thought to participate in ECM formation, cytokine secretion, cell signal transduction, transcriptional regulation, and apoptosis (Xie et al., 2004). In an effort to develop chemically defined culture conditions for hES cells a number of feeder- and serum-free protocols have been developed (Avery et al., 2006 and references within). Basic fibroblast growth factor (bFGF/FGF2) has been added to all these media formulations, and it appears that FGF plays a similar role in maintaining hES cells as LIF in maintaining mES cells. A number of other factors expressed and secreted by MEFs and/or provided by serum have been identified and these include activin A, TGFβ, BMP antagonists noggin and gremlin (Xu et al., 2005), Wnt (Sato et al., 2004; Dravid et al., 2005), and neurotrophins (Pyle et al., 2006). Although these multiple factors appear to play a role in maintaining self-renewal, preventing differentiation, and

Embryonic and Extraembryonic Stem Cells

1C.3.13 Current Protocols in Stem Cell Biology

Supplement 3

promoting cell survival, it is still unclear if they are sufficient to prolong the pluripotent state of hES cells and allow for their clonal expansion. Although mES cells can be grown conveniently in a medium supplemented with LIF, it is recommended that stock mES cells be grown on feeder layers to prevent culture deterioration and to keep cells pluripotent (Evans, 2004). At this stage, prolonged culture of hES cells can best be achieved on feeder cell layers, because despite extensive research, factors affecting their long-term maintenance are still not well defined. The development of totally defined, animal product-free culture conditions that support long-term growth of diploid hESCs is an important goal for research and clinical applications of these cells. The concern over xeno-contamination from MEFs and FBS led to the development of a number of “humanized” culture systems (Mallon et al., 2006 and references within; Lei et al., 2007 and references within). These include derivation of feeder cell lines from adult and fetal human tissues and from differentiated hESC cultures, substitution of feeders with human and mouse-derived extracellular matrices, and replacement of FBS with serum replacement (SR). Derivation and propagation of hESCs in chemically defined culture media and on human-derived substrate has also been reported (Fletcher et al., 2006; Ludwig et al., 2006). However, it has to be pointed out that feeder-free culture can lead to chromosomal abnormalities (Draper et al., 2004), while prolonged culture in currently available chemically defined, xeno-free media does not seem to maintain undifferentiated growth of hESCs (Rajala et al., 2007).

Critical Parameters and Troubleshooting

Isolation and Preparation of MEFs

It is important that all aspects of media and reagents preparation are carried out aseptically. If possible, most sterile culture media and solutions should be obtained from commercial suppliers. However, if solutions are prepared in house, tissue-culture-grade reagents and water should be used. These solutions should be sterilized by autoclaving or filtration through a 0.2-µm pore filter. However, it is important to stress that filtration will not remove mycoplasmas or viruses. If contamination with these microorganisms is suspected, solutions should be discarded. All glassware used for tissue culture reagents should be kept separate from general laboratory glassware.

The glassware should be washed and rinsed thoroughly to remove all traces of detergent and should be heat-sterilized, including washing and sterilization (2 hr at 180◦ C), rather than autoclaved. The quality of fibroblast feeder cells is crucial for the maintenance of undifferentiated ES cells. MEFs must be isolated and propagated aseptically, passaged at appropriate density, and efficiently inactivated to prevent their growth while maintaining metabolic activity. In most laboratories, antibiotics are routinely added to all culture media and can mask lowlevel contamination resulting from poor aseptic techniques. It is recommended that, after initial isolation and expansion, antibiotics be omitted from the culture media. All new batches should also be routinely tested for the presence of mycoplasma. The infection with this microorganism does not cause medium turbidity and can persist undetected for a considerable length of time spreading throughout all cultures in the laboratory. All contaminated cultures should be discarded immediately. MEFs cultured in vitro can undergo ∼20 divisions before entering senescence (Karatza and Shall, 1984). It is recommended that MEFs are not expanded beyond passage 5. With time, the number of dividing cells diminishes and the cell morphology changes from tightly packed pavement-like to large, fried egg-like or elongated, stringy appearance. Such fibroblasts will produce inferior, nonsupportive feeders and should be discarded. Occasionally, cells will grow very slowly after initial dissociation of fetuses. This can result from inadequate tissue mincing and reduced trypsin activity or from excessive digestion with trypsin. This can also occur when very small fetuses (from a large litter) or fetuses younger than 12.5 dpc are used. Although such slow-growing cultures can be rescued by low splitting ratio or pooling cells from a number of flasks, they tend to produce feeders that will not support hES cell cultures. It is recommended that a new batch of MEFs be isolated. Both, mitomycin C treatment and γirradiation are effective in mitotic inactivation of MEFs. However, occasionally some cells escape inactivation and foci of growing fibroblast colonies appear in feeder layers. In such cases fresh mytomicin C solution should be prepared and/or time of treatment increased. If γ-irradiation was used, the source of radiation should be calibrated and irradiation titration performed, to assure effective and controlled inactivation.

1C.3.14 Supplement 3

Current Protocols in Stem Cell Biology

Plating density of inactivated MEFs depends on particular culture conditions and application and will affect the feeder cell morphology. At the higher density, recommended for mouse ES cells, the feeder’s morphology will resemble that of growing fibroblasts while at the lower cell density, required for human ES cells, the feeder cells will be spread out, larger, and more circular. Since MEFs are primary cell lines, they have a limited lifespan and fresh batches must be prepared routinely and frequently. This might result in a batch-to-batch variation and necessitates routine testing. Generally, mouse ES cells do not seem to be affected by these variations; however, it is important that new batches are tested for their efficiency to support human ES cells. It is recommended that hES cells are passaged on a new batch for a minimum of 3 to 4 weeks to determine the quality of MEFs. It is not uncommon to see an increased degree of differentiation after the first passage. However, over time hES cells should become adapted to new feeders. MEFs supporting the growth and maintenance of ES cells have been isolated from a number of different strains of mice and the choice of the strain will depend on availability and particular requirements (e.g., drug resistance necessary for genetic manipulation of ES cells). The advantage of using an outbred or F1 strain is that a larger litter is obtained. On the other hand, some researchers claim that MEFs isolated from an inbred strain such as 129Sv will produce fibroblast cell layers that provide better support of hES cell culture (Reubinoff et al., 2000).

Anticipated Results Isolation of MEFs is a straightforward and efficient procedure. From an average litter of 6 to 10 fetuses a stock of 6 to 12 early passage vials can be frozen (passage 1; 107 cells per vial). It is expected that an 80% to 90% confluent 75-cm2 flask of MEFs will yield ∼8 to 10 × 106 cells. Each batch should be equivalent in its ability to maintain pluripotent mouse ES cells. However, different batches of MEFs can differ in their ability to support human ES cells and some batches may be nonsupportive. Therefore, it is crucial, that all newly isolated MEFs are tested for a minimum of 3 to 4 passages of human ES cells.

Time Considerations It is recommended that time-mated female mice are ordered in advance, as not all matings result in pregnancy. When ordering from an Current Protocols in Stem Cell Biology

outside supplier, day 12.5 to 14.5 confirmed pregnant females should be requested. When females are mated in house, some time for acclimation before mating might be required. The time necessary for dissection of fetuses will depend on the researcher’s experience and the number of fetuses used. Approximately 1.5 to 2 hr should be allocated for processing a single litter of 6 to 10 fetuses. After plating, cultures should reach 80% to 90% confluency within 1 to 3 days. Subsequently, MEFs should be passaged every 2 to 4 days. Passaging or freezing of MEFs will take from 20 min to 1 hr, depending on the number of flasks. Thawing MEFs will take 20 to 30 min. Mitomycin C treatment and feeder layer plating will require ∼3.5 to 4 hr (including 2 to 3 hr incubation). Approximately 1 hr should be allocated for the mycoplasma testing.

Literature Cited Avery, S., Inniss, K., and Moore, H. 2006. The regulation of self-renewal in human embryonic stem cells. Stem Cells Dev. 15:729-740. Daheron, L., Opitz, S.L., Zaehres, H., Lensch, W.M., Andrews, P.W., Itskovitz-Eldor, J., and Daley, G.Q. 2004. LIF/STAT3 signaling fails to maintain self-renewal of human embryonic stem cells. Stem Cells 22:770-778. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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. Donovan, J.D. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Draper, J.S., Smith, K., Gokhalel, P., Moore, H.D., Maltby, E., Johnson, J., Meisner, L., Zwaka, T.P., Thomson, J.A., and Andrews, P.W. 2004. Recurrent gain of chromosomes 17q and 12 in cultured human embryonic stem cells. Nat. Biotechnol. 22:53-54. Dravid, G., Ye, Z., Hammond, H., Chen, G., Pyle, A.D., Donovan, P.J., Yu, X., and Cheng, L. 2005. Defining role of Wnt/β-catenin signaling in the survival, proliferation, and self-renewal of human embryonic stem cells. Stem Cells 23:14891501. Evans, M. 2004. Isolation and maintenance of murine embryonic stem cells. In Handbook of Stem Cells, Vol. 1: Embryonic Stem Cells (R. Lanza, J. Gearhart, B. Hogan, D. Melton, R. Pedersen, J. Thomson, and M. West, eds.) pp. 413-417. Elsevier Academic Press, Amsterdam. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Ezashi, T., Das, P., and Roberts, R.M. 2005. Low O2 tensions and the prevention of differentiation of hES cells. Proc. Nat. Acad. Sci. U.S.A. 2005:4783-4788.

Embryonic and Extraembryonic Stem Cells

1C.3.15 Supplement 3

Fletcher, J.M., Ferrier, P.M., Gardner, J.O., Harkness, L., Dhanjal, S., Serhal, P., Harper, J., Delhanty, J., Brownstein, D.G., Prasad, Y.R., Lebkowski, J., Mandalam, R., Wilmut, I., and De Dousa, P.A. 2006. Variations in humanized and defined culture conditions supporting derivation of new human embryonic stem cell lines. Cloning Stem Cells 8:319-334. Forsyth, N.R., Musio, A., Vezzoni, P., Simpson, A.H., Noble, B.S., and McWhir, J. 2006. Physiologic oxygen enhances human embryonic stem cell clonal recovery and reduces chromosomal abnormalities. Cloning Stem Cells 8:16-23. Gardner, L.R. 2004. Isolation and maintenance of murine embryonic stem cells. In Handbook of Stem Cells, Vol. 1: Embryonic Stem Cells (R. Lanza, J. Gearhart, B. Hogan, D. Melton, R. Pedersen, J. Thomson, and M. West, eds.) pp. 15-26. Elsevier Academic Press, Amsterdam.

Ogawa, K., Nishinakamurab, R., Iwamatsua, Y., Shimosatoa, D., and Niwa, H. 2006. Synergistic action of Wnt and LIF in maintaining pluripotency of mouse ES cells. Biochem. Biophys. Res. Commun. 343:159-166. Ogiso, Y., Kume, A., Nishimune, Y., and Matsushiro, A. 1982. Reversible and irreversible stages in the transition of cell surface marker during the differentiation of pluripotent teratocarcinoma cell induced with retinoic acid. Exp. Cell Res. 137:365-372. Parrinello, S., Samper, E., Krtolica, A., Goldstein, J., Melov, S., and Campisi, J. 2003. Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nature Cell Biol. 5:741747.

Karatza, C. and Shall, S. 1984. The reproductive potential of normal mouse embryo fibroblasts during culture in vitro. J. Cell Sci. 66:401409.

Pera, M.F., Andrade, J., Houssami, S., Reubinoff, B., Trounson, A., Stanley, E.G., Ward-van Oostwaard, D., and Mummery, C. 2004. J. Cell Sci. 117:1269-1280.

Lei, T., Jacob, S., Ajil-Zaraa, I., Dubuisson, J-B., Irion, O., Jaconi, M., and Feki, A. 2007. Xenofree derivation and culture of human embryonic stem cells: Current status, problems and challenges. Cell Res. 17:682-688.

Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18.

Lim, J.W.E. and Bodnar, A. 2002. Proteome analysis of conditioned medium from mouse embryonic fibroblast feeder layers which support the growth of human embryonic stem cells. Proteomics 2:1187-1203. Ludwig, T.E., Levenstein, M.E., Jones, J.M., Berggren, W.T., Mitchen, E.R., Frane, J.L., Crandall, L.J., Daigh, C.A., Conard, K.R., Piekarczyk, M.S., Llanas, R.A., and Thomson, J.A. 2006. Derivation of human embryonic stem cells in defined conditions. Nat. Biotechnol. 24:185-187. Mallon, B.S., Park, K.Y., Chen, K.G., Hamilton, R.S., and McKay, R.D.G. 2006. Toward xenofree culture of human embryonic stem cells. Int. J. Biochem. Cell Biol. 38:1063-1075. Martin, G.R. 1981. Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc. Nat. Acad. Sci. U.S.A. 78:76347638. Martin, G.R. and Evans, M.J. 1975. Differentiation of clonal lines of teratocarcinoma cells: formation of embryoid bodies in vitro. Proc. Nat. Acad. Sci. U.S.A. 72:1441-1445. Matsuda, T., Nakamura, T., Nakao, K, Arai, T., Katsuki, M, Heike, T., and Yokota, T. 1999. STAT3 activation is sufficient to maintain an undifferentiated state of mouse embryonic stem cells. EMBO J. 18:4261-4269.

Isolation and Preparation of MEFs

Ogawa, K., Matsui, H., Ohtsuka, S., and Niwa, H. 2004. A novel mechanism for regulating clonal propagation of mouse ES cells. Genes Cells 9:471-477.

Niva, H., Burdon, T., Chambers, I., and Smith, A. 1998. Self-renewal of pluripotent embryonic stem cells is mediated via activation of STAT3. Genes Dev. 12:2048-2060.

Ponchio, L. Duma, L., Oliviero, B., Gibelli, N., Pedrazzoli, P., and Robustelli della Cuna, G. 2000. Mitomycin C as an alternative to irradiation to inhibit the feeder layer growth in longterm culture assays. Cytotherapy 2:281-286. Puck, T.T. and Marcus, P.I. 1955. A rapid method for viable cell titration and clone production with HeLa cells in tissue culture: the use of xirradiated cells to supply conditioning factors. Proc. Nat. Acad. Sci. U.S.A. 4:432-437. Pyle, A.D., Lock, L.F., and Donovan, P.J. 2006. Neurotrophins mediate human embryonic stem cell survival. Nat. Biotechnol. 24:344-350. Rajala, K., Hakala, H., Panula, S., Aivio, S., Pihlajamaki, H., Suuronen, R., Hovatta, O., and Skottman, H. 2007. Testing of nine different xeno-free culture media for human embryonic stem cell cultures. Hum. Reprod. 22:1231-1238. Rathjen, P.D., Toth, S., Willis, A., Heath, J.K., and Smith, A.G. 1990. Differentiation inhibiting activity is produced in matrix-associated and diffusible forms that are generated by alternate promoter usage. Cell 62:1105-1114. Reubinoff, B., Pera, M.F., Fong, C., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399-404. Roy, A., Krzykwa, E., Lemieux, R., and Neron, S. 2001. Increased efficiency of gamma-irradiated versus mitomycin C-treated feeder cells for the expansion of normal human cells in long-term cultures. J. Hematother. Stem Cell Res. 10:873880. Sato, N., Meijer, L., Skaltsounis, L., Greengard, P., and Brivanlou, A.H. 2004. Maintenance of

1C.3.16 Supplement 3

Current Protocols in Stem Cell Biology

pluripotency in human and mouse embryonic stem cells through activation of Wnt signaling by a pharmacological GSK-3-specific inhibitor. Nat. Med. 10:55-63. Smith, A.G., Heath, J.K., Donaldson, D.D., Wong, G.G., Moreau, J., Stahl, M., and Rogers, D. 1988. Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336:688-690. Suemori, H. and Nakatsuji, N. 1987. Establishment of the embryo-derived stem (ES) cell lines from mouse blastocysts: Effects of the feeder cell layer. Dev. Growth Differ. 29:133-139. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 228:1145-1147. Wang, F., Thirumangalathu, S., and Loeken, M.R. 2006. Establishment of new mouse embryonic stem cell lines is improved by physiological glucose and oxygen. Cloning Stem Cells 8:108116. Ware, L.M. and Axelrad, A.A. 1972. Inherited resistance to N- and B-tropic murine leukemia viruses in vitro: Evidence that congenic mouse strains SIM and SIM.R differ at the Fv-1 locus. Virology 50:339-348. Williams, R.L., Hilton, D.J., Pease, S., Willson, T.A., Stewart, C.L., Gearing, D.P, Wagner, E.F., Metcalf, D., Nicola, N.A., and Gough, N.M. 1988. Myeloid leukaemia inhibitory factor maintains the developmental potential of embryonic stem cells. Nature 336:684-687. Wobus, A.M., Holzhausen, J¨akel, P., and Sch¨oneich, J. 1984. Characterization of a pluripotent stem cell line derived from a mouse embryo. Exp. Cell Res. 152:212-219. Xie, C.Q., Lin, G., Luo, K.L., Luo, S.W., and Lu, G.X. 2004. Newly expressed proteins of mouse

embryonic fibroblasts irradiated to be inactive. Biochem. Biophys. Res. Commun. 315:581-588. Xie, C.Q., Lin, G., Yuan, D., Wang, J., Liu, T.C., and Lu, G.X. 2005. Proliferative feeder cells support prolonged expansion of human embryonic stem cells. Cell Biol. Int. 29:623-628. Xu, R.H., Chen, X., Li, D.S., Li, R., Addicks, G.C., Glennon, C., Zwaka, T.P., and Thomson, J.A. 2002. BMP4 initiates human embryonic stem cell differentiation to trophoblast. Nat. Biotechnol. 20:1261-1264. Xu, R.H., Peck, R.M., Li, D.S., Feng, X., Ludwig, T., and Thomson, J.A. 2005. Basic FGF and suppression of BMP signaling sustain undifferentiated proliferation of human ES cells. Nat. Methods 2:185-190. Ying, Q.L., Nichols, J., Chambers, J., and Smith, A. 2003. BMP induction of Id proteins suppresses differentiation and sustains embryonic stem cell self-renewal in collaboration with STAT3. Cell 115:281-292.

Key References Lanza, R., ed. 2004. Handbook of Stem Cells, Vol. 1: Embryonic Stem Cells (R. Lanza, J. Gearhart, B. Hogan, D. Melton, R. Pedersen, J. Thomson, and M. West, eds.) pp. 15-26. Elsevier Academic Press, Amsterdam. Collection of articles by world’s experts in the field of embryonic stem cell research, providing background information and methods on various aspects of ES manipulation. Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. 2002. Manipulating the Mouse Embryo: A Laboratory Manual. 3rd Ed. Cold Spring Harbor Press, New York. A comprehensive manual describing all aspects of early mouse embryo culture and manipulation including isolation and propagation of mouse embryonic stem cells.

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1C.3.17 Current Protocols in Stem Cell Biology

Supplement 3

Culture of Mouse Embryonic Stem Cells 1

1

Gabi Tremml, Matthew Singer, and Richard Malavarca 1

UNIT 1C.4

1

Millipore Corporation, Bioscience Division, Billerica, Massachusetts

ABSTRACT In this unit standard culture conditions for mouse embryonic stem cells (mESCs) on primary murine embryonic fibroblast (PMEF or MEF) monolayers, culture conditions without MEF for feeder-independent mESCs, and culture conditions in chemically defined media for both feeder-independent mESCs and feeder-dependent mESCs are described. For expansion of an mESC line, it is crucial that cells maintain their undifferentiated state and their self-renewal capacity, and that they remain karyotypically normal, all of which are necessary for successful chimerization of the germ line upon blastocyst injection. Derivation and culture conditions for the original mESCs have been described (notably Robertson, 1987; Smith, 1991; Nagy et al., 2003), however, as there are more and more mESC lines available, it becomes evident that culture conditions are cell-line specific to some extent, and there is a constant demand for culturing details for mESC lines derived C 2008 by from different mouse strains. Curr. Protoc. Stem Cell Biol. 5:1C.4.1-1C.4.19.
John Wiley & Sons, Inc. Keywords: mouse embryonic stem cell (mESC) r murine embryonic fibroblasts (MEF) r fetal bovine serum (FBS) r leukemia inhibitory factor (LIF) r trypsin/EDTA r chemically defined media r ESGRO complete r Accutase r enzyme-free dissociation solution

INTRODUCTION The derivation and culture of mESCs in vitro 20 years ago has led to enormous possibilities in the field of mouse genetics. If cultured correctly, mESCs retain the potential to differentiate into all three germ layers and can successfully chimerize the germ line upon blastocyst injection. Optimal culture implies that mESCs maintain their undifferentiated state and self-renewal capacity, and that no gross karyotypic abnormalities are introduced. Many different mESC lines derived from various mouse strains are available to date that can be successfully used in mouse modeling, and it has become evident that the choice of the genetic background of these lines is important for certain disease models. It has also become evident that different mESC lines can require different culture conditions. Traditionally, most mESCs are cultured on primary murine embryonic fibroblast (PMEF or MEF) monolayers, however, more recently, other culture conditions have been established. Here, culture conditions for mESCs on MEF monolayers (see Basic Protocol), culture conditions without MEF for feeder-independent mESCs (see Alternate Protocol 1), and culture conditions in chemically defined media for both feeder-independent mESCs and feeder-dependent mESCs (see Alternate Protocols 2 and 3, respectively) are described. Protocols for serum testing (see Support Protocol 1), cryopreservation for mESCs (see Support Protocol 2), and counting mESCs (see Support Protocol 3) are also provided.

STANDARD MOUSE EMBRYONIC STEM CELL CULTURE ON MEFs Two key signaling molecules are involved in maintaining self-renewal of mESCs, leukemia inhibitory factor (LIF; Suda et al., 1987; Niwa et al., 1998) and bone morphogenic protein 4 (BMP-4; Ying et al., 2003). Most cell lines derived from the 129 and C57Bl/6 mouse strains need LIF provided from two sources, (1) supplemented in

Current Protocols in Stem Cell Biology 1C.4.1-1C.4.19 Published online April 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c04s5 C 2008 John Wiley & Sons, Inc. Copyright


BASIC PROTOCOL

Embryonic and Extraembryonic Stem Cells

1C.4.1 Supplement 5

the culture medium and (2) secreted from the mitotically inactivated MEFs, generally plated the day before on the culture flask or dish as a feeder monolayer. MEFs can be inactivated with mytomycin C treatment or MEFs can be irradiated; the isolation and treatment procedure is described elsewhere in detail (Conner, 2000; UNIT 1C.3). The MEF monolayer provides both identified and unidentified factors, as well as cell-cell contact necessary for optimal growth. Expansion of feeder-dependent mESCs also uses the addition of serum, usually fetal bovine serum (FBS), into the medium, the quality of which is crucial for optimal culturing conditions. If purchasing serum that is not pre-qualified for use with mESCs, testing of serum prior to its use with mESCs is described in Support Protocol 1. Here, optimal culture conditions for 129Sv/Ev and C57Bl/6J (Bruce 4) cells, both of which are originally feeder-dependent mESC lines, is described. This protocol is useful for all other 129 and C57Bl/6 strains; it is also useful for mESCs derived from other strains, however, minor adjustments may have to be made to optimize culture for a particular mESC line. During subculture of mESCs, the medium should be replaced daily, and generally, cells should be split every other day. A vigorously growing mESC line may be split 1:5, while a slow growing line may be split at 1:2. One vial of frozen mESCs usually contains 2.5 × 106 cells, and can be seeded onto one 100-mm dish. NOTE: All the protocols using mESC and MEF cells require either a laminar flow hood or a safety cabinet (allowing for sterile tissue culture) and a 37◦ C water bath. NOTE: All media used for tissue culture should be pre-warmed for at least 30 min in a 37◦ C water bath. NOTE: Aseptic working techniques are required, and all solutions and materials coming into contact with live cells must be sterile. Use 70% isopropanol and sterile wipes to clean the hood area before handling cells and the pre-warmed bottles of media and wear latex gloves. NOTE: All incubations are carried out in a 37◦ C, 5% CO2 humidified incubator unless stated otherwise. NOTE: The use of materials and cell lines listed here is consistent with Biohazard level 2.

Materials 0.1% gelatin (Millipore cat. no. SF008) Hygromycin-resistant MEFs (Millipore cat. no. PMEF-H) Neomycin-resistant MEFs (Millipore cat. no. PMEF-N) MEF medium (see recipe), prewarmed mESC medium (see recipe), prewarmed 129SvEv – ES cells (Millipore cat. no. CMTI-1) C57BL6/J – ES cells (Millipore cat. no. CMTI-2) D-PBS without Ca2+ and Mg2+ (CMF-DPBS; Millipore cat. no. BSS-1006-A) D-PBS (Millipore cat. no. SM-2002-C) 0.05% trypsin/EDTA (Millipore cat. no. SM-2002-C)

Culture of Mouse Embryonic Stem Cells

100-mm tissue culture dishes (25-cm2 or 75-cm2 , Falcon) 37◦ C water bath 15- and 50-ml tubes (Falcon) Swing-out tissue culture centrifuge High-quality microscope with 10× and 20× oculars

1C.4.2 Supplement 5

Current Protocols in Stem Cell Biology

Table 1C.4.1 MEF Cell Densities and Medium Volumes for Plating

Dish/flask Volume of medium size (ml)

Growth area (cm2 )

Number of MEF cells/well or flask

75-cm2

12

75

3.75 × 106

25-cm2

6

25

1.25 × 106

100-mm

10

56

2.8 × 106

60-mm

5

21

1 × 106

6-well

4

9.6

4.75 × 105

12-well

2

4

2 × 105

24-well

1

2

1 × 105

96-well

0.1

0.32

1.5 × 104

Prepare MEF dishes (day 1) 1. Estimate how many dishes will be needed (including at least one 1:2 to 1:5 split, and following experiments). Generally, one vial of mESCs contains 2.5 × 106 cells, and can be plated on one 100-mm dish or equivalent.

2. Pre-coat culture dishes or flasks with 0.1% gelatin solution. Use just enough volume to cover dish surface (e.g., 5 to 10 ml for a 100-mm dish). Incubate at least 30 min at room temperature. 3. Estimate how many vials of MEFs will be needed for the number of dishes determined in step 1. One vial of MEF cells contains 5–7 × 106 MEFs. MEF cell densities for a confluent feeder layer and MEF medium volumes are described in Table 1C.4.1.

4. Rapidly thaw vials of MEF cells in a 37◦ C water bath until just the last bit of ice is thawed and then allow further thawing as the cells are aseptically transferred to 10or 50-ml tubes containing prewarmed MEF medium. For one vial of MEF cells, use 10 ml of MEF medium. Multiple vials can be combined into the same tube and can be from different MEF preparations.

5. Centrifuge 5 min at 168 × g, 4◦ C. 6. Aspirate the medium and resuspend the cell pellet in the appropriate volume of MEF medium. Remove the gelatin solution from the dishes or flasks and immediately plate MEFs onto them without letting the gelatin dry. Incubate the MEFs overnight. Plate density according to Table 1C.4.1 and use the volumes indicated. For calculation of cell density, use the cell number indicated on the MEF cryovials. These cell numbers are adjusted to allow for some cell death if stored correctly, according to the manufacturer’s directions. Therefore, on the following day, some cell death can be observed, but the plated cell number will be correct. Keep in mind that the cell numbers on the MEF cryovials are approximate numbers, but if a confirmation of the number is required, after prolonged storage, for example, MEF cells should be resuspended and a viable count should be performed according to Support Protocol 3.

7. On the next morning, observe settled feeder monolayer under a high-quality microscope and make sure that the layer is confluent (i.e., the entire plastic area is covered with feeder cells). Prepared dishes with confluent monolayers of mitotically inactive MEFs can be kept in the incubator for up to 1 week.

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Thaw mESCs (day 2) 8. Prepare a 15-ml tube with 10 ml of prewarmed mESC medium. 9. Thaw one vial of mESCs rapidly in a 37◦ C water bath. 10. Thaw the vials until just before the last bit of ice is thawed and then allow thawing to continue as the cells are transferred. Aseptically transfer mESCs into 10- or 50-ml tubes with 10 ml of prewarmed mESC medium. 11. Centrifuge 5 min at 168 × g, 4◦ C. Aspirate or decant supernatant. 12. Resuspend mESCs with 10 ml of fresh mESC medium into a single-cell suspension by pipetting up and down.

Figure 1C.4.1 129 SvEv and C57Bl6 mESC cultures in standard media on MEF feeder monolayers. (A) 129SvEv mESCs in trypsin/EDTA after 10 min. (B) MEF feeder monolayer. (C) 129 SvEv mESCs 1 day after thawing. (D) 129SvEv 2 days in culture, just before passage. (E) C57Bl/6 mESC 2 days in culture, alkaline phosphatase staining. (F) 129SvEv mESC 6 days in culture with overgrowth and differentiation. Culture of Mouse Embryonic Stem Cells

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Current Protocols in Stem Cell Biology

Figure 1C.4.2 Culture of mESCs in absence of MEF feeder monolayers. (A–D) culture in standard medium. (E–H) culture in chemically defined ESGRO Complete clonal-grade medium. (A, B, E, H) E14 mESCs the day after passage. (C, D, G, H) R1 mESCs the day after passage. (B, D, F, H) Alkaline phosphatase staining.

13. Remove MEF medium from the culture plates prepared with feeder monolayers (from step 7). 14. Gently mix mESCs again, and transfer the entire contents onto the culture dish or flask with feeder monolayers. 15. Incubate overnight. Embryonic and Extraembryonic Stem Cells

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Figure 1C.4.3 Sequential adaptation of C57Bl/6 mESCs to the chemically defined feederfree/serum-free ESGRO Complete clonal-grade medium. (A) mESCs in enzyme-free dissociation solution after 10 min. (B) mESCs on gelatin-coated dishes, at passage 2 in the appropriate medium mixture; note that MEFs are still present. (C) mESCs at passage 4 in the appropriate medium mixture. (D) mESCs at passage 6 in ESGRO Complete clonal-grade medium. (E) mESCs passage 7 in ESGRO Complete clonal-grade medium; note that MEFs are depleted. (F) Alkaline phosphatase staining of passage 7 mESCs.

Feed cultures 16. Examine mESCs under the microscope, there should be small bright colonies visible (see Fig. 1C.4.1). 17. Decide whether mESCs are ready to be passaged. If so, proceed to step 20. Generally, mESCs are not ready the day after the thaw. Occasionally, even if the colonies are still small, it can be anticipated that the growth will be fast over the next night, and the dish will be over-confluent the next day. Alternatively, the dish may be 80% confluent already. In this case, the mESCs should be passaged, proceed to day 4. Culture of Mouse Embryonic Stem Cells

18. If cells are not ready to be passaged, remove mESC medium and replace with fresh prewarmed medium. 19. Incubate overnight.

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Passage mESCs 20. Examine mESCs under the microscope, and decide whether the cells should be passaged. Colonies are bright, big, and individual cells are difficult to see. The colonies do not touch each other. See Figure 1C.4.1D.

21. Have culture dishes ready containing a confluent monolayer of MEFs. 22. For passage, remove the MEF medium and wash dish or flask with CMF-DPBS, using ∼10 ml or more for a 100-mm dish. 23. Remove CMF-DPBS and add just enough 0.05% trypsin/EDTA to cover the cells. For a 100-mm dish, use ∼3 ml.

24. Incubate at 37◦ C and observe dissociation of mESCs under the microscope. Incubate only until mESCs begin to detach from the culture dish surface (5 to 10 min). 25. Add 5 ml of mESC medium and pipet up and down, without causing foaming or bubbles, the entire contents of the dish. It is important to obtain a single-cell solution, as aggregates of mESCs tend to differentiate.

26. Transfer the dissociated mESC solution into a 15-ml tube and centrifuge 5 min at 168 × g, 4◦ C. 27. Resuspend mESCs in 10 ml of mESC medium, and pipet up and down to assure a single-cell solution. Count cells (see Support Protocol 3) and transfer 2.5 × 106 (usually 1/2 to 1/5 of a 100-mm dish) mESCs into previously prepared gelatin-coated 100-mm culture dishes with confluent monolayers of mitotically inactivated MEFs. Do not use dishes with monolayers of MEFs older than 1 week.

SERUM TESTING For the novice, buying serum that is ES-qualified is recommended. For the experienced mESC culturist, the following protocol describes serum testing for pre-qualification. Tests on three to five serum lots are recommended. ES qualification guarantees that cells of either the 129SvEv or the C57Bl/6J (Bruce4) background can be cultured. Therefore, the qualified serum will generally work for mESCs of other backgrounds.

SUPPORT PROTOCOL 1

Materials mESCs in culture (see Basic Protocol) Samples of three to five FBS serum lots Additional reagents and equipment for culturing mESCs (see Basic Protocol) 1. Culture mESC for at least two passages as described in the Basic Protocol. Anticipate how many 6-well plates are needed for the test, and prepare feeder plates accordingly (one serum lot test will require 8 wells, including a control). 2. Prepare mESC medium with 10%, 15%, and 30% of the test FBS, and keep a control medium for mESC with the current serum lot. Media can be prepared the day before the plating of mESCs.

3. Passage mESCs into gelatin-coated wells prepared with MEF monolayers; seed mESCs in each medium in duplicate. Seed 1000 to 1500 cells per well in prepared mESC medium with the test serum and include a control for mESC medium with the current serum.

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4. On the next day, remove media and replace with fresh media. 5. Incubate for 5 days, and observe growth every day. 6. Compare morphology of the mESC colonies to the control. Watch for differentiation, colony size, colony number, and general appearance of the well. An FBS lot is acceptable when differentiation is minimal at 10% and 15% FBS concentration and little cell death is observed in the 30% condition. Compare degree of differentiation, colony size and colony number, and cell death to the control medium with pre-qualified FBS. Tests on three to five serum lots are recommended for the qualification of one serum. ALTERNATE PROTOCOL 1

mESC CULTURE WITHOUT MEFs IN STANDARD MEDIUM Here the culture of mESCs without MEF feeder layers in standard medium is described. There are some mESC lines available that tolerate growth only on gelatin-coated culture dishes without differentiation. Many of the 129 lines and all C57Bl/6 lines will differentiate rapidly when depleted of MEFs in standard serum-containing mESC medium, despite the presence of LIF. However, some 129 lines including R1, E14Tg2, and D3 have been found to sustain vigorous growth in absence of MEF feeder layers. This protocol is very similar to the Basic Protocol for mESCs cultured on MEFs. There is no need for plating MEFs the day before; dishes and flasks are coated with 0.1% gelatin at room temperature for 30 min prior to plating the mESCs. The mESC medium is replaced every day and mESCs will need to be passaged every other day. Daily microscopic examination of colonies and cell morphologies is required. Feeder-independent mESCs grow more vigorously, and cells can be passaged at 1:5 to 1:10. This protocol is very similar to that of the feeder-dependent mESCs, but it will take into account the absence of MEFs and the growing vigor of the R1, D3, and E14 mESC lines under feederindependent conditions.

Additional Materials (also see Basic Protocol) Standard mESC medium (see recipe), prewarmed mESC lines: E14Tg2, R1, D3 Thaw mESCs (day 1) 1. Coat a 25-cm2 flask with 0.1% gelatin, incubate at least 30 min at room temperature and remove before plating mESCs. Do not allow the gelatin to dry out by quickly plating mESCs after the 30-min incubation. 2. Prepare a 15-ml tube with 10 ml prewarmed standard mESC medium. 3. Rapidly thaw one vial of mESCs (one vial usually contains 2.5 × 106 cells) in a 37◦ C water bath. 4. Aseptically transfer mESCs into the 15-ml tube with 10 ml of prewarmed standard mESC medium and gently mix. 5. Centrifuge mESCs 5 min at 168 × g in a swing-out rotor centrifuge at 4◦ C. 6. Discard supernatant and resuspend mESCs in 10 ml of prewarmed standard mESC medium. Triturate to assure a single-cell suspension. 7. Transfer cells to gelatin-coated flask. Place cells in incubator overnight. Culture of Mouse Embryonic Stem Cells

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Feed cultures 8. Examine mESCs under the microscope. Morphology is slightly different than that of mESCs cultured on MEFs. Colonies appear flatter and less shiny.

9. Determine whether passaging is required. If passaging is required, proceed to step 12. Typically, cells need 1 day for growth, but occasionally they might be ready for passaging.

10. Replace medium with 10 to 15 ml prewarmed standard mESC medium for a 25-cm2 tissue culture flask. 11. Place cells back into the incubator overnight.

Passage cells (day 3) 12. Examine mESCs under the microscope and determine whether passaging is required. Colonies should be 70% to 90% confluent.

13. For passage, remove medium and add 10 ml CMF-DPBS. Rock gently for 10 sec and remove CMF-DPBS. Ensure that all CMF-DPBS is removed. 14. Repeat step 13. 15. Add just enough trypsin/EDTA to cover the culture dish surface (1.5 ml for 25-cm2 flask) and place in the incubator for 5 min or until dissociated clumps of mESCs are visible under the microscope. 16. Add 10 ml of prewarmed standard mESC medium, triturate, and transfer the entire contents into a 15-ml tube. 17. Centrifuge 5 min at 168 × g, 4◦ C. 18. Resuspend mESCs in 10 ml standard mESC medium.Count cells and add 1 × 106 cells (usually 1/5 to 1/10 of a 25-cm2 flask) to a fresh gelatin-coated flask. 19. Place cells back into incubator overnight.

mESC CULTURE USING CHEMICALLY DEFINED MEDIA: SEQUENTIAL ADAPTATION

ALTERNATE PROTOCOL 2

Sometimes controlled and reproducible expansion of mESCs is preferred without the variable effects of each FBS lot. Controlled expansion implies that the mESC medium is chemically defined and no serum is present. The obvious major advantages are that there is no need for MEFs to be prepared the day before, and there is no need to prescreen various lots of FBS. Here, the adaptation and culturing conditions of feeder-free E14Tg2a and R1 mESCs and the feeder-dependent C57Bl/6 and 129SvEv from serumcontaining medium to serum- and feeder-free conditions in chemically defined medium are described. The sequential adaptation is generally used for feeder-dependent mESC lines, while the direct adaptation protocol (see Alternate Protocol 3) is used for feeder-free mESCs. The Alternate Protocols 2 and 3 are primarily recommended for the experienced mESC culturist. This protocol is recommended for a first-time adaptation of a mESC line from conditions where serum and/or feeders are used. It uses mixtures of mESC (i.e., serumcontaining) medium and the ESGRO Complete clonal-grade medium as described in Table 1C.4.2. Cells are passaged using trypsin/EDTA when regular serum-containing medium is present and Accutase or enzyme-free dissociation solution when 100% serumfree ESGRO clonal-grade medium is used. Feeders are naturally depleted as mESCs are passaged. Once mESCs are adapted, cells are passaged every 2 to 4 days and medium is replaced with fresh medium every 2 days. Current Protocols in Stem Cell Biology

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Table 1C.4.2 Sequential Adaptation to Chemically Defined Medium

Medium

% mESC medium % Clonal-grade medium

Passage number 1

2

3

4

5

6

100

80

50

20

0

0

0

20

50

80

100

100

Additional Materials (also see Basic Protocol) mESC medium (see recipe) ESGRO Complete clonal-grade medium (Millipore cat. no. SF001) Accutase (Millipore cat. no. SF006) Enzyme-free dissociation solution (Millipore cat. no. SF009) ESGRO Complete basal medium (Millipore cat. no. SF002) Prepare cells 1. Culture mESCs on feeder layers using mESC medium until confluent in a 25-cm2 flask or culture dish for at least one to two passages adhering to the Basic Protocol. 2. On the day of beginning the adaptation, pre-coat 25-cm2 flasks with 0.1% gelatin solution for at least 30 min at room temperature. Remove gelatin before plating mESCs; but do not allow gelatin to dry out.

Progressively adapt cells to chemically defined medium 3. Prepare medium mixture containing 80% of mESC medium and 20% ESGRO Complete clonal-grade medium. Passage mESCs using trypsin/EDTA and adhering to the Basic Protocol for splitting mESCs. 4. Seed mESCs on gelatin-coated 25-cm2 flasks in the medium mixture from step 3 at a density of 1 × 106 per 25-cm2 flask. Place flasks overnight in incubator. 5. On the following day, there will still be plenty of MEFs left among the cells that have adhered. Determine whether it is necessary to passage cells; alternatively, observe growth for another day. 6. When flasks are subconfluent (60% to 70% confluent), prepare medium containing 50% mESC medium and 50% ESGRO Complete clonal-grade medium and prepare fresh gelatin-coated flasks. 7. Repeat steps 2 to 4 using a 50:50 mix of mESC medium/ESGRO Complete clonalgrade medium. 8. On the following day, there will be significantly fewer MEFs present. Observe growth under the microscope. Usually, an additional day of incubation is required, but it may be necessary to passage cells at this point. 9. When flasks are subconfluent (60% to 70% confluent), prepare medium containing 20% mESC medium and 80% ESGRO Complete clonal-grade medium and prepare fresh gelatin-coated flasks. 10. Repeat steps 2 to 4 using the 20:80 mix of mESC medium/ESGRO Complete clonalgrade medium. Culture of Mouse Embryonic Stem Cells

11. On the following day, observe that few or no MEFs should be present among the adherent cells. Incubate for an additional 1 to 2 days and observe colony formation— colonies look flatter and the nucleus becomes visible. Do not grow flask more than subconfluent.

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Plate cells in chemically defined medium 12. When flasks are subconfluent, prepare fresh gelatin-coated flasks. 13. Add 10 ml CMF-DPBS, gently rock flask for 10 sec, and then remove CMF-DPBS. Do not decant to prevent introduction of contamination. 14. To dissociate cells, add 1.5 ml Accutase or enzyme-free dissociation solution per 25-cm2 flask. Incubate at 37◦ C and allow cells to detach (no more than 5 to 10 min, often as little as 3 min with Accutase). Tap flask to dissociate colonies. If necessary, triturate to obtain a single-cell solution. Either Accutase or enzyme-free dissociation solution can be used; Accutase may be slightly more aggressive, and the enzyme-free dissociation solution may take longer.

15. Add 10 ml of ESGRO Complete basal medium, mix, and centrifuge 5 min at 168 × g, 4◦ C. 16. Remove supernatant and repeat step 15. 17. Resuspend cell pellet in 10 ml of 100% ESGRO Complete clonal-grade medium and seed at a concentration of 2 × 106 cells into a gelatin-coated 25-cm2 flask. Observe growth for 1 to 2 days in a 37◦ C incubator. Colonies are flatter, some cell death and differentiation will become apparent and will continue as cells are passaged further. Always keep cells subconfluent (60% to 70% confluence).

18. Repeat passage into ESGRO Complete clonal-grade medium for two to three more passages until mESCs are considered to be adapted to serum- and feeder-free culture conditions.

mESC CULTURE USING CHEMICALLY DEFINED MEDIA: DIRECT ADAPTATION

ALTERNATE PROTOCOL 3

The following protocol is applicable for adapting feeder-independent mouse ES cells to serum-free cell culture conditions.

Additional Materials (also see Basic Protocol and Alternate Protocol 2) Feeder-free mESC cultures (see Alternate Protocol 2) 1. Grow feeder-free mouse ES cells to 60% confluence in mESC medium in a 25-cm2 flask without feeders. 2. Change medium 24 hr prior to seeding into ESGRO complete clonal-grade medium. 3. Pre-coat 25-cm2 flasks with 0.1% gelatin solution (see Basic Protocol). 4. Wash cells once with 10 ml CMF-DPBS and remove all CMF-DPBS (see Basic Protocol). To dissociate cells, add 1.5 ml Accutase. Incubate at 37◦ C until the cells begin to detach (3 to 10 min). It is important not to use standard trypsin/EDTA solutions, as“balling” of ES cells will occur after the cells are re-plated.

5. Gently tap and add 5 ml of ESGRO Complete basal medium, mix gently, and centrifuge 5 min at 168 × g, 4◦ C. 6. Remove supernatant and resuspend pellet in 5 ml ESGRO Complete clonal-grade medium. Count cells. 7. Plate 1 × 106 cells into a pre-coated 25-cm2 flask containing 10 ml pre-warmed ESGRO Complete clonal-grade medium.

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8. Observe cell growth over the next 2 to 3 days in a 37◦ C incubator. Some cell death may be observed and some differentiation may be visible. However, ES cell colonies will continue to grow and may appear to be flatter than those on feeder cells with a distinct nuclear and cytoplasmic morphology.

9. When mESCs are ∼60% confluent, passage again into a new gelatin-coated 25-cm2 flask containing 10 ml ESGRO Complete clonal-grade medium. Always be careful not to allow mESCs in serum-free medium become over-confluent as they will begin to differentiate. SUPPORT PROTOCOL 2

CRYOPRESERVATION OF mESCs As a rule, mESCs are cryopreserved slowly and gradually to −80◦ C (at −1◦ C/min) before storage in liquid nitrogen to prevent water crystals from forming inside the cells, thereby preventing increased cell death when thawing. Thawing should be done quickly in a 37◦ C water bath. Cryopreservation (Nagy et al., 2003) can be similarly done with all mESCs independent of which culture protocol has been used. When working with cells grown in serum-free conditions, serum-free freezing solutions should be used. When freezing mESCs keep a record of passage number, date, and location of the cryovials. Generally, mESCs are frozen at a concentration of 2–5 × 106 cells/ml. This translates into two to four cryovials per 100-mm dish, assuming that the dish can culture up to 1 × 107 cells (two vials if a concentration of 5 × 106 /ml is used and four vials if a concentration of 2 × 106 /ml is used). The following conditions can be applied to all mESCs.

Materials mESCs DPBS without Ca2+ and Mg2+ (CMF-DPBS; Millipore cat. no. BSS-1006-A) 0.05% trypsin/EDTA (Millipore cat. no. SM-2002-C), Accutase (Millipore cat. no. SF006), or enzyme-free dissociation solution (Millipore cat. no. SF009) mESC medium (see recipe) or ESGRO complete basal medium (see Alternate Protocols 2 and 3) 2× ES-qualified freezing medium (Millipore cat. no. ES002D) Liquid nitrogen tank ESGRO complete freezing medium (serum-free; Millipore cat. no. SF005) Cryovials (inside thread; Nunc) 15-ml tubes Freezing containers (Nunc) Styrofoam 37◦ C water bath CAUTION: Freezing solutions contain DMSO, see MSDS for handling, storage, and disposal. 1. Estimate the number of cryovials needed and label cryovials with strain, passage, number of cells, and the date, as well as any other relevant information. 2. Wash mESCs with 10 ml CMF-DPBS. Gently rock dish or flask for 10 sec at room temperature and remove all DPBS. 3. Add trypsin/EDTA (see Basic Protocol and Alternate Protocol 1), Accutase, or enzyme-free dissociation solution (see Alternate Protocols 2 and 3). Culture of Mouse Embryonic Stem Cells

Add 4 ml per 100-mm dish to cover all cells.

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4. Incubate 5 to 10 min at 37◦ C, or the time needed to dissociate colonies (see Basic Protocol and Alternate Protocols 1 to 3). Pipet up and down to obtain a single-cell solution. 5. Transfer to a 15-ml tube, dilute with 10 ml mESC medium or ESGRO complete basal medium and centrifuge 5 min at 168 × g, 4◦ C. 6. Resuspend cells in 10 ml fresh mESC medium or ESGRO complete basal medium. Remove 10 µl of the suspension and count cells (see Support Protocol 3). Centrifuge 5 min at 168 × g, 4◦ C. 7. Calculate the volume of freezing medium and how many cryovials are needed to obtain 2.5 × 106 cells/0.5 ml. If serum containing mESC medium has been used (such as in the Basic Protocol and Alternate Protocol 2), use 2× ES-qualified freezing medium and proceed to step 8. For one 100-mm dish containing 1 × 107 mESCs cultured in serum-containing medium, four cryovials and 1 ml of 2× freezing medium and 1 ml mESC serum-containing medium is required. If serum-free chemically defined medium is used (such as in Alternate Protocols 2 and 3), then the ESGRO complete freezing medium is required. For one 100-mm dish containing 1× 107 mESCs cultured in ESGRO Clonal medium, four cryovials and 2 ml of ESGRO complete freezing medium is required. Spin cells down and resuspend in 2 ml of ESGRO complete feezing medium. Dispense 0.5 ml into each cryovial. For one 100-mm dish containing 1 × 107 mESCs, four vials and 2 ml of 1× freezing solution are required.

8. Resuspend mESCs to 1/2 of the final volume with mESC medium, then add an equal volume of the cold 2× ES-qualified freezing medium to obtain a 1× solution. Mix gently and add 0.5 ml into each cryovial. 9. Place cryovials in a Styrofoam box, and place into a −80◦ C freezer. 10. On the next day, transfer vials to a liquid nitrogen tank for extended storage. mESCs can be kept up to 4 weeks at −80◦ C, longer storage requires liquid nitrogen storage.

11. For thawing, late in the day prepare a 15-ml tube with 10 ml prewarmed mESC medium. Have the appropriate culture dish ready, with prepared MEFs if necessary. Alternatively, thaw as described in Basic Protocol.

12. Place vial with mESCs into a 37◦ C water bath for a rapid thaw until just before all the ice has melted. Freezing medium volume should not exceed 0.5 ml. Cell number should be known.

13. Aseptically transfer the mESCs into the 10 ml medium and gently mix. 14. Replace the medium on the culture dish with the contents of the 15-ml tube. Incubate overnight. 15. On the next morning, replace medium with 10 ml prewarmed mESC medium.

COUNTING mESCs mESCs should be counted and viability assessed when thawing and passaging cells.

SUPPORT PROTOCOL 3

Materials mESCs to be counted Appropriate mESC medium

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0.4% (w/v) Trypan blue (Invitrogen cat. no. 15250-061) Hemacytometer 1. Resuspend mESCs in 10 ml of appropriate medium into a single-cell suspension. 2. Mix gently and remove 10 µl of cell suspension. 3. Mix with 10 µl of 0.4% Trypan blue. 4. Load 10 µl of the mix into a hemacytometer. 5. Count live cells in one to nine quadrants, dead cells will be blue. Distinguish between MEF and mESCs. Live mESCs are round and shiny with a smooth surface, MEFs have a rough surface and are not perfectly round.

6. Count 15 to 50 cells in each quadrant. If counts are outside 15 to 50 cells, then the result will be inaccurate. If >50 cells are present, dilute cell suspension and reload the hemacytometer. If <15 cells are counted, re-centrifuge cells and resuspend mESCs in a smaller volume. 7. Calculate the number of cells (count in one quadrant gives the number of cells × 104 /ml). Total cell count = average cell count in one quadrant × 104 × suspension volume, ml (10) × dilution (2).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

MEF medium For a 570 ml total volume, add: 500 ml ES cell–qualified DMEM (Millipore cat. no. SLM-220-M) 50 ml FBS ES-qualified serum (Millipore cat. no. ES-009-B) 5 ml 100× nucleosides (Millipore cat. no. ES-008-D) 5 ml penicillin-streptomycin (Millipore cat. no. TMS-AB2-C) 5 ml non-essential amino acids (Millipore cat. no. TMS-001-C) 5 ml L-glutamine solution (Millipore cat. no. TMS-002-C) Store freshly prepared medium up to 4 weeks at 4◦ C mESC medium For a 605 ml total volume, add: 500 ml ES cell–qualified DMEM (Millipore cat. no. SLM-220-M) 75 ml FBS ES-qualified serum (Millipore cat. no. ES-009-B) 5 ml 100× nucleosides (Millipore cat. no. ES-008-D) 5 ml penicillin-streptomycin (Millipore cat. no. TMS-AB2-C) 5 ml non-essential amino acids (Millipore cat. no. TMS-001-C) 5 ml L-glutamine solution (Millipore cat. no. TMS-002-C) 5 ml 2-mercaptoethanol (Millipore cat. no. ES-007-E) 5 ml LIF (ESGRO 105 U/ml; Millipore cat. no. ESG1107) Store freshly prepared medium for 3 weeks at 4◦ C Culture of Mouse Embryonic Stem Cells

Dilute 1 ml ESGRO (107 U) in total 100 ml of complete mESC medium then filter sterilize. Store in 5-ml aliquots at −20◦ C for extended storage.

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Standard mESC medium For a 580 ml total volume, add: 500 ml ES cell–qualified DMEM (Millipore cat. no. SLM-220-M) 50 ml FBS ES-qualified serum (Millipore cat. no. ES-009-B) 5 ml 100× nucleosides (Millipore cat. no. ES-008-D) 5 ml penicillin-streptomycin (Millipore cat. no. TMS-AB2-C) 5 ml non-essential amino acids (Millipore cat. no. TMS-001-C) 5 ml L-glutamine solution (Millipore cat. no. TMS-002-C) 5 ml 2-mercaptoethanol (Millipore cat. no. ES-007-E) 5 ml LIF (ESGRO 105 U/ml; Millipore, cat. no. ESG1107) Store freshly prepared medium for 3 weeks at 4◦ C Dilute 1 ml ESGRO (107 U) in total 100 ml of complete mESC medium then filter sterilize. Store in 5-ml aliquots at −20◦ C for extended storage.

COMMENTARY Background Information Culturing mESCs with MEFs Pluripotent mESCs are isolated from the inner cell mass of a blastocyst embryo (Evans and Kaufman, 1981; Martin, 1981; Bradley et al., 1984; Robertson et al., 1987), and much research addresses the question of how pluripotency is being maintained in mESCs (reviewed by Niwa, 2007). mESCs are extensively used for genetic manipulation using homologous recombination (Wurst and Joyner, 1993; Nagy et al., 2003) and differentiation studies (Smith, 1991). Many of the original mESCs are robust cell lines, and will remain pluripotent over many passages. Culturing mESCs on MEF feeders is accepted widely as the best method for expansion, as it provides the best assurance that the cells will remain pluripotent. However, for some of the original mESC lines, STO or SNL (STO lines transfected with a NeoR and LIF expressing vector) feeder cell lines have been used with good results. Other more recently developed cell lines, especially C57Bl/6, require MEFs to be present as a feeder layer, i.e., they are not amenable to growth on STO or SNL monolayers or gelatin-coated dishes with standard media. mESC culture in standard medium without MEFs Although best results are achieved when mESCs are cultured on primary MEFs, there are a few unique cell lines that tolerate culture conditions without MEFs and are just cultured on gelatin-coated dishes (Ward et al., 2002). In these cases, the mESCs do not require secreted factors, nor cell-cell contact with MEFs, and the LIF provided in the culture medium keeps the mESCs in the

undifferentiated state. Such classical feederindependent mESC lines are E14Tg2a, derived from the 129/Ola mouse strain (Hooper et al., 1987; Nichols et al., 1990), R1, derived from the progeny of strains 129×1/SvJ x 129S1 (Nagy et al., 1993), and D3, derived from the 129S2/SvPas strain (Doetschman et al., 1985). Addition of compounds into standard media may improve culture conditions not only for the classical cell lines but may also improve maintenance of pluripotency for many different mESC lines adapted and cultured in absence of MEFs. Studies in mESCs revealed that glycogen synthase kinase-3β inhibitors supplemented into the culture medium maintain pluripotency and self-renewal through the Wnt pathway in the absence of MEFs (Sato et al., 2004; Umehara et al., 2007). Also, more recently, a chemical screen identified a small molecule SC1, also called pluripotin, that maintains self-renewal of mESCs when added to standard media in the absence of MEFs, and OG2 mESCs were successfully expanded in SC1-supplemented medium (Chen et al., 2006). Culture of mESCs in chemically defined medium Traditionally, culture of mESCs involves the use of serum and LIF in addition to MEF feeder layers. Ying et al. (2003) have shown that the use of BMP4 together with LIF replaces the need for serum and feeder cells for ES self-renewal. mESCs cultured in the presence of BMP4 and LIF on gelatin-coated plastic dishes or flasks will retain their ability to differentiate along multiple lineages, produce chimeras, and even contribute to the germ line when injected into blastocyst hosts. This serum- and feeder-free system has been used to develop the ESGRO Complete clonal medium.

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This medium has been used to derive pluripotent mESC lines from the 129 mouse strain, a strain that is permissive for relatively effortless mESC derivation from the inner cell mass. Moreover, mESCs from the non-permissive C56Bl/6 strain have been derived under serumand feeder-free conditions; these cells have contributed to germ line transmission when injected into Balb/C morulae (Nichols and Ying, 2006).

Critical Parameters and Troubleshooting

Culture of Mouse Embryonic Stem Cells

Culturing mESCs with MEFs mESCs have the capacity to self-renew and to remain pluripotent, which allows for indefinite growth in vitro under perfect culture conditions. However, continued passaging can have an effect on genetic content, integrity, and differentiation status of mESCs. Therefore, one critical parameter is, when starting with mESC work, that vials of low passages are provided. Passage 1 refers to the first single colony isolated from the inner cell mass, and passage number refers to the number of times a culture has been split into new culture vessels. There are only a very limited number of vials from passage 6 of any mESC line available; these are stored typically for expansion to passage 9 or more. Generally, a passage 11 vial will be good for expansion for one or more experiments, such as electroporation or differentiation studies. However, as researchers prefer lower-numbered passages, it is recommended that during expansion vials of high-quality mESCs be cryopreserved for storage and future experiments. It is important to acknowledge that a small percentage of mESCs do acquire karyotypic abnormalities over time, and cells with certain chromosomal aberrations may gain a selective growth advantage over normal mESCs. This is particularly true for Trisomy 8 in mESCs (Liu et al., 1997). Therefore, it is always recommended prior to an expensive and time-consuming experiment, especially the generation of chimeric mice with blastocyst injections, that mESCs be karyotyped. While karyotyping with G-banding will only show gross chromosomal abnormalities, it will evaluate cells for their stability in general, and a good karyotype will have a normal chromosome count in >80% of the cells. A normal karyotype in <60% of the cells generally is associated with significantly decreased success for germ line transmission (Longo et al., 1997; Suzuki et al., 1997).

Another critical parameter is to judge correctly when and how to passage a culture dish. A dish never should be >80% confluent, colonies should not be touching or fusing with each other as they will start differentiating, and colonies should not be too sparse, as cells will undergo crisis. For a vigorously growing mESC line, this means that cells will need to be passaged every 2 days at optimal cell densities, while for a slow-growing mESC line, passaging every 3 days may be appropriate. Optimal cell densities can vary; while 1 × 106 mESCs or less will be appropriate for seeding with a vigorously growing mESC line in a 25-cm2 flask, a slow growing mESC line may need to be seeded at 2 × 106 mESCs. Counting cells with a hemacytometer during a passage is inevitable for the novice culturist or when handling a new mESC line, but with experience and knowledge of the growth properties of a particular mESC line, cell densities can be estimated under the microscope. Appearance of mESC colonies needs to be microscopically examined daily; colonies should show shiny white and closed borders. Morphological changes of colonies can include opening of borders and loss of shiny appearance, both of which are signs of differentiation and are usually a result of incorrect culturing. Drastic morphological changes of cells include the appearance of nucleus resulting from a decrease in the nuclear-to-cytoplasm ratio. Absence of differentiation can be assessed by morphological criteria and markers of pluripotency. Alkaline phosphatase staining (Thomson et al., 1998), Oct4 antibody staining, and stagespecific embryonic antigen SSEA-1 staining (Henderson et al., 2002; Muramatsu and Muramatsu, 2004) will show little or no differentiation in >90% of the colonies of a highquality mESC line. All the assessments above combined will allow for a general prediction of pluripotency and therefore the ability of the mESCs to populate the germ line. One of the difficulties in culturing mESCs are changes in the growing pattern and sudden differentiation visible by morphology. At times, the material, lot numbers of solutions, supplier, or handling of the materials can be the origin of these changes. Therefore, it is crucial that all the expiration dates supplied by the manufacturer are observed, media discarded after recommended time, and critical material such as serum be tested prior to use. Certain sera might be perfectly usable for a

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particular mESC line while cells of another line may tend to differentiate when that same serum lot is used. Similarly, certain mESC lines are not sensitive to the number of MEFs on the dish, and variable densities of monolayers can be used, while other lines need a denser MEF monolayer, and will tend to differentiate when the number of MEFs is lower. Even change of tissue culture plastic can affect mESC growth. The authors found that both 129 and C57Bl/6 grow best on Nunc plates. Lastly, mESCs should be tested for pathogens, such as mycoplasma and other difficult-to-detect microorganisms, as the presence of these can influence germ line transmission rate. mESC culture in standard medium without MEFs Feeder-independent mESCs grow rapidly and mESC medium can become acidic, thereby changing the color of the color indicator to orange or yellow. This is usually a sign that the mESCs are overcrowded and cells will undergo extensive differentiation. It is crucial that an optimal cell density is maintained (seeding of 1 × 106 in a 25-cm2 flask); too high or too low a density will lead to excessive differentiation and cell death. Sometimes, it is necessary to passage cells every 24 hr to ensure maintenance of pluripotency. Additionally, great care should be taken that cells are plated as a single-cell suspension, and that the suspension is evenly dispersed in the culture dish or flask. Cell aggregates and uneven plating will lead to differentiating cell clumps and areas of massive differentiation. Culture of mESCs in chemically defined medium During adaptation and culturing of mESCs in serum- and feeder-free clonal-grade medium, the morphology of the colonies and of single cells changes. The colonies become considerably flatter and the nucleus and cytoplasm become visible. Even with optimal culturing conditions, increased cell death might be observed, especially during adaptation but also during general expansion and passaging. However, if culture conditions are kept optimal according to the protocol described, mESCs will remain pluripotent. It is critical when passing cells in Accutase or enzyme-free dissociation solution that incubation time is no longer than 5 to 10 min or the least amount of time required for disso-

ciation; with Accutase, the incubation can be halted when cells are first observed to be detaching from the culture dish surface (in as little as 3min). It is equally important that mESCs are resuspended into a single-cell suspension, otherwise the cells tend to clump together to form floating colonies. Compared to regular trypsin/EDTA solutions, Accutase or enzymefree dissociation solution requires more rigorous resuspension. Another critical parameter is to keep the mESCs sub-confluent (∼60% to 70%), as mES cells tend to differentiate in more confluent conditions. Keep in mind that colonies look flatter than when grown on MEFs, and the nucleus and cytoplasm may become visible. Even though cells look morphologically different if grown in feeder- and serum-free conditions, they should remain pluripotent if cultured correctly, as it has been shown for E14, C57Bl/6, and 129 mESC lines (Ward et al., 2002, using Invitrogen’s Serum Replacement Knockout SR cat. no. 10829-018; Ying et al., 2003; Nichols and Ying, 2006, using ESGRO Complete) When seeding mESCs, as a general rule to stay within 1–2 × 106 cells per 25-cm2 flask, it is necessary to count cells during passages. When mESCs are too dense, they will start differentiating rapidly. If mESCs are not dense enough, they will undergo crisis, resulting in increased cell death. There are some mESC lines with which successful adaptation and culturing in serumand feeder-free conditions remains difficult. Cryopreservation Cell death after thawing mESCs from cryopreservation is inevitable. To assess cell death, perform a live count with Trypan blue and seed cells according to the number of live cells present. When thawing mESCs where the thawed cells are diluted into medium, make sure that the freezing medium containing DMSO does not exceed 5% to 10% when diluted into the mESC medium. Since DMSO can induce the differentiation of ES cells, changing the medium first thing on the following morning will minimize the effects of residual DMSO. Alternatively, the safest method to prevent DMSO-associated effects is to wash mESCs before plating. Do not freeze more than five to ten vials at one time, as extended time in DMSO at room temperature will increase cell death.

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Anticipated Results Culturing mESCs on MEFs Starting out with 1 × 10 6 mESCs, a passage of 1:3 every other day, theoretically, should yield cells up to 2.7 × 107 after 7 days of culture under optimally controlled conditions. mESC culture in standard medium without MEFs Starting out with 1 × 106 mESCs, a passage of 1:5 every other day theoretically should yield 1.25 × 108 mESCs after 7 days of culture under optimal culture conditions. Keep in mind that the morphology of mESCs cultured in absence of MEFs in standard medium always shows some feeder-like cells on the edges of colonies. This is expected and should not influence the outcome of the experiment. Culture of mESCs in chemically defined medium Doubling time varies among different mESC lines in chemically defined media. The E14 and other robust cell lines will continue with vigorous growth, as one might be able to passage them up to 1:5 every other day. Other cell lines like 129 SvEv and C57Bl/6 will have a slower doubling time and might be passaged only 1:2 to 1:3 every other day or even every third day. Cryopreservation Cell death should not be >30% if correct storage conditions were applied; there will be floating dead cells the day after the thaw.

Time Considerations Culturing mESCs with MEFs When starting mESC culture, always take into account that MEFs have to be plated the day before thawing mESCs. Once mESCs are thawed, cells need to grow for at least one or two passages before they can be cryopreserved again; this becomes logistically important, especially if there is weekend work involved. Hands-on work might be only 1 to 2 hr a day when expanding one vial of mESCs, but it is important that cultures be observed every day.

Culture of Mouse Embryonic Stem Cells

Culture of mESCs in chemically defined medium Adaptation to chemically defined culture conditions requires ∼2 weeks of mESCs in culture, after that, cells can be safely cyropreserved. Thawing out adapted mESCs requires at least 2 passages before cyropreservation.

Daily microscopic examination is required, as well as hands-on work for ∼1 to 2 hr per day. Cryopreservation Cryopreservation of mESCs is done at the end of an experiment and the procedure itself takes just 30 min of hands-on time for one to five vials of mESCs.

Literature Cited Bradley, A., Evans, M., Kaufman, M.H., and Robertson, E. 1984. Formation of germ-line chimaeras from embryo-derived teratocarcinoma cell lines. Nature 309:255-256. Chen, S., Do, J.T., Zhang, Q., Yao, S., Yan, F., Peters, E., Sch¨oler, H.R., Schultz, P.G., and Ding, S. 2006. Self-renewal of embryonic stem cells by a small molecule. Proc. Natl. Acad. Sci. U.S.A. 103:17266-17271. Conner, D.A. 2000. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Mol. Biol. 51:23.2.1-23.2.7. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Henderson, J.K., Draper, J.S., Baillie, H.S., Fishel, S., Thomson, J.A., Moore, H., and Andrews, P.W. 2002. Preimplantation human embryos and embryonic stem cells show comparable expression of stage-specific embryonic antigens. Stem Cells 20:329-337. Hooper, M.L., Hardy, K., Handyside, A., Hunter, S., and Monk, M. 1987. HPRT deficient (LeschNyhan) mouse embryos derived from germline colonization by cultured cells. Nature 236: 292295. Liu, X., Wu, H., Loring, J., Hormuzdi, S., Disteche, C.M., Bornstein, P., and Jaenisch, R. 1997. Trisomy eight in ES cells is a common potential problem in gene targeting and interferes with germ line transmission. Dev. Dyn. 209:85-91. Longo, L., Bygrave, A., Grossveld, F.G., and Pandolfi, P.P. 1997. The chromosome make-up of mouse embryonic stem cells is predictive of somatic and germ cell chimaerism. Transgenic Res. 6:321-328. Martin, G.R. 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. 78:76347638. Muramatsu, T. and Muramatsu, H. 2004. Carbohydrate antigens expressed on stem cells and early embryonic cells. Glycoconjugate J. 21:41-45. Nagy, A., Rossant, J., Nagy, R., AbramowNewerly, W., and Roder, J. 1993. Derivation of completely cell culture–derived mice from

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early-passage embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 90:8424-8428. Nagy, A., Gertsensten, M., Vintersten, K., and Behringer, R. 2003. Manipulating the Mouse Embryo: A Laboratory Manual. 3rd edition, Cold Spring Harbor Press, Cold Spring Harbor, NY. Nichols, J. and Ying, Q.I. 2006. Derivation and propagation of embryonic stem cells in serumand feeder-free culture. Methods Mol. Biol. 329:91-98. Nichols, J., Evans, E.P., and Smith, A.G. 1990. Establishment of germ-line competent embryonic stem (ES) cells using differentiation inhibiting activity. Development 110:1341-1348. Niwa, H. 2007. How is pluripotency determined and maintained? Development 134:635-646. Niwa, H., Burdon, T., Chambers, I., and Smith, A. 1998. Self-renewal of pluripotent embryonic stem cells is mediated via activation of STAT3. Genes Dev. 12:2048-2060. Robertson, E., Bradley, A., Kuehn, M., and Evans, M. 1986. Germ-line transmission of genes introduced into cultured pluripotential cells by retroviral vector. Nature 323:445-448. Robertson, E.J. 1987. Embryo-derived stem cell lines. In Teratocarcinomas and Embryonic Stem Cells: A Practical Approach (E.J. Robertson, ed.) pp. 71-112. IRL Press, Oxford. Sato, N., Meijer, L., Skaltsounis, L., Greengard, P., and Brivanlou, A.H. 2004. Maintenance of pluripotency in human and mouse embryonic stem cells through activation of Wnt signaling by a pharmacolocial GSK-3-specific inhibitor. Nature Medicine 1:55-63. Smith, A.G. 1991. Culture and differentiation of embryonic stem cells. J. Tiss. Cult. Meth. 13:8994. Suda, Y., Suzuki, M., Ikawa, Y., and Aizawa, S. 1987. Mouse embryonic stem cells exhibit indefinite proliferative potential. J. Cell Physio. 133:197-201. Suzuki, H., Kamada, N., Ueda, O., Jishage, K., Kurihara, H., Terauchi, Y., Azuma, S., Kadowaki, T., Kodama, T., Yazaki, Y., and

Toyoda, Y. 1997. Germ-line contribution of embryonic stem cells in chimeric mice: Influence of karyotype and in vitro differentiation ability. Exp. Anim. Tokyo 46:17-23. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Umehara, H., Kimura, T., Ohtsuka, S., Nakamura, T., and Kitajima, K. 2007. Efficient derivation of embryonic stem cells by inhibition of glycogen synthase kinase-3. Stem Cells: published online Jul 19. Ward, C.M., Stern, P., Willington, M.A., and Flenniken, A.M. 2002. Efficient germline transmission of mouse embryonic stem cells grown in synthetic serum in the absence of a fibroblast feeder layer. Laboratory Investigations 82:12,P1765-1767. Wurst, W. and Joyner, A. 1993. Production of targeted embryonic stem cell clones. In Gene Targeting: A Practical Approach (A. Joyner, ed.) pp. 33-62. IRL Press at Oxford University Press. Ying, Q.I., Nichols, J., Chambers, I., and Smith, A. 2003. BMP induction of Id proteins suppresses differentiation and sustains embryonic stem cell self-renewal in collaboration with STAT3. Cell 115:281-292.

Internet Resources http://mailman.ic.ac.uk/mailman/listinfo/ transgenic-list This Web site hosts an electronic discussion forum. The list members include active researchers in transgenesis from novice to the experts in the field. Major keywords cover a wide spectrum of disciplines: homologous recombination, targeted mutagenesis, inducible expression, ES cells, microinjection, mouse genetics, and animal husbandry. http://www.informatics.jax.org/mgihome/lists/ lists.shtml This Web site hosts an electronic discussion forum for topics in mouse genetics.

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Preparation of Autogenic Human Feeder Cells for Growth of Human Embryonic Stem Cells

UNIT 1C.5

Rodolfo Gonzalez,1, 2 Jeanne F. Loring,3 and Evan Y. Snyder2 1

University of California, San Diego, La Jolla, California Burnham Institute for Medical Research, La Jolla, California 3 Scripps Research Institute, La Jolla, California 2

ABSTRACT Human embryonic stem cells (hESCs) that are currently distributed under NIH guidelines, as well as many of those that are not on the NIH registry, have been derived and maintained in coculture with growth-arrested mouse embryonic fibroblasts (MEFs). Using this mouse support system may compromise the therapeutic potential of these hESCs because of the risk of transmitting xenopathogens. Alternatively, to reduce this risk, methods to culture undifferentiated hESCs on autologous hESC-derived human feeder layers have now been developed. This feeder cell system derived from hESCs successfully prolongs growth of undifferentiated hESCs and eliminates risk factors and concerns about using xenogeneic or unknown allogeneic feeders. In this unit, we provide the necessary protocols for an autogeneic human feeder system that efficiently supports hESC growth and maintenance C 2008 by John Wiley of pluripotency. Curr. Protoc. Stem Cell Biol. 4:1C.5.1-1C.5.15.
& Sons, Inc. Keywords: human embryonic stem cells r fibroblast-like r autologous r feeders

INTRODUCTION Culturing human embryonic stem cells (hESCs) requires a significant commitment of time and resources. It takes weeks to establish a culture, and the cultures require daily attention. Once hESC cultures are established, they can, with skill and the methods described below, be kept in continuous culture for years. hESC lines were originally derived using very similar culture medium and conditions as those developed for the derivation and culture of mouse ESC lines. However, these methods were suboptimal for hESCs, and have evolved considerably in the years since hESC lines were derived. Compared with mouse ESCs, hESCs are very difficult to culture—they grow slowly, and most importantly, since we have no equivalent assay for germline competence, we cannot assume that the cells that we have in our culture dishes are either stable or pluripotent. This makes it far more critical to assay the cells frequently, using characterization methods such as karyotyping, immunocytochemistry, gene expression analysis, and fluorescence activated cell sorting (FACS).

Tips for successfully culturing hESCs 1. Feed cells every day, except for 1 or 2 days following passage. 2. Examine the cultures every day using 4× and 10× phase-contrast microscopy. This will allow you to become familiar with the morphologies of undifferentiated and differentiated cells and colonies. Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1C.5.1-1C.5.15 Published online March 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c05s4 C 2008 John Wiley & Sons, Inc. Copyright


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3. When they are cultured on hESC-derived feeder layers, some hESC lines tend to undergo spontaneous differentiation in the centers of the colonies. When passaging, take care to avoid passaging these differentiated “center” hESC to the new culture. 4. Most hESC lines double every 31 to 35 hr. 5. Store medium at 4◦ C and discard any unused medium after 10 days. Best results are achieved when medium is prepared in small batches once a week. This unit presents methods for preparation of stocks of hESC-derived feeders (Basic Protocol), passaging by mechanical dissection (Support Protocol 1), passaging by enzymatic dissection (Support Protocol 2), and growth of hESCs in MEF-conditioned medium (Support Protocol 3). NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. BASIC PROTOCOL

PREPARATION OF STOCK hESC-DERIVED FEEDER CELLS This protocol describes the necessary steps to obtain an unlimited supply of hESC-derived feeder cells for hESC growth. The hESC-derived feeders are fibroblast-like cells (Fig. 1C.5.2). The rate of growth of these cells is the same as for undifferentiated hESCs. These cells have been characterized by us using microarray, immunohistochemistry, and flow cytometry. They are positive for the following cell-surface proteins: PDGFRα, SEMA5A, F2RL2, endoglin, ALCAM, and CD44. The feeder layer plays a complex role in helping to maintain hESCs in an undifferentiated state. The feeder layer must be healthy and rapidly dividing prior to inactivation in order to provide the best substrate for the growth of the hESCs. Following inactivation, feeder layer cells remain adequate for the culture of hESCs for 5 to 7 days. In order to keep the feeder layer healthy, it is advisable to change the medium on the inactivated cells every 3 days. Always observe the feeder layer under the microscope prior to using for the culture of hESCs, in order to confirm that the feeder layer is still intact and the cells have not begun to deteriorate. Since hESCs are usually passaged every 5 to 7 days, the feeder layer can start to deteriorate before the hESCs are ready to passage if “old” feeder layer dishes are used. For best results, inactivate the feeders the day before passaging the hESCs. Inactivation of the feeder cells is accomplished by either irradiation or treatment with mitomycin C. Inactivated feeder cells are usually plated on gelatin-coated dishes to aid in their attachment.

Materials

Autogenic Human Feeder Cells for hESCs

hESC cultures grown under feeder-free conditions (Support Protocol 3) in 6-well plates MEF-conditioned medium (Support Protocol 3) Calcium- and magnesium-free Dulbecco’s phosphate-buffered saline (CMF-DPBS; e.g., Invitrogen or Sigma) Trypsin/EDTA: 0.05% (w/v) trypsin/0.53 mM EDTA (Invitrogen) Feeder medium (see recipe) hESC medium (see recipe) Mitomycin C medium (see recipe) Freezing medium (90 % v/v FBS/10% v/v DMSO), ice cold Liquid nitrogen

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Low-power dissecting microscope Pipettor with 20-µl sterile filter tip, or 23-G needle 15-ml conical tubes Centrifuge Gelatin-coated 75-cm2 culture flasks (and other appropriately sized vessels as needed): add 0.1% (w/v) gelatin to dishes/flasks at 1 ml/cm2 and leave 1 hr to overnight at room temperature; just before plating the cells remove gelatin and rinse with CMF-DPBS γ irradiator (Nordion Gammacell 40 Exactor, or equivalent low-dose 137 Cs irradiator for both cells and whole animals with central dose rate of ∼1.10 Gy/min (∼110 rad/min). 1-ml cryovials Isopropanol freezing container Additional reagents and equipment for growing hESCs under feeder-free growth conditions (Support Protocol 3) NOTE: Volumes are described for one well of a 6-well plate or 35-mm tissue culture dish. They should be adjusted for other sizes of tissue culture vessels based on the relative surface area.

Isolate hESC-derived feeders 1. Grow hESCs in feeder-free growth culture conditions in MEF-conditioned medium (see Support Protocol 3) in 6-well plates for 7 days. By day 7 of culture, hESC-derived feeder-like cells at the periphery of hESC will be readily visible (Fig. 1C.5.1A). IMPORTANT NOTE: Some colonies may not have any visible, feeder-like cells.

2. Under a low-power dissecting microscope (in a horizontal flow hood), dissect all of the undifferentiated hESC colonies from each well of the plate using a pipettor with a 20-µl sterile filter tip attached, or a sterile 23-G needle.

Figure 1C.5.1 nification.

An undifferentiated hESC (WA09) colony growing on mouse feeders, 10× mag-

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3. Break up each colony by moving the tip around and across each colony in a spiral motion, being careful not to disrupt the peripheral hESC-derived feeders. Since the colonies are large by day 7 of hESC culture, it is relatively easy to see individual colonies on the plate and, with practice, one can quickly dissociate an entire plate in less than 20 min.

4. After all of the colonies are dissected (Fig. 1C.5.1B), use a 5-ml pipet to transfer the culture medium containing the dissected colonies to a 15-ml conical tube. Rinse the plate two times with 3 ml CMF-DPBS to remove any remaining hESC clusters and transfer to the 15-ml tube with the culture medium. The cells derived from the dissected colonies are hESCs, not feeder cells, and will be replated separately from the feeder cells, which are those obtained by trypsinization in steps 5 to 6. The cells in the tube are passaged as hESCs, as described in Support Protocol 3, and they will continue to grow as pluripotent, undifferentiated hESCs.

5. Add 1 ml of trypsin/EDTA per well of the 6-well plate from which the hESC colonies have been dissected. Incubate at 37◦ C for 3 to 5 min until all the hESC-derived feeders lift up from the wells. 6. Add 1 ml of feeder medium per well and combine all of the hESC-derived feeders by transferring all of the cell suspension from each well into a 15-ml conical tube. Centrifuge 5 min at 350 × g, 4◦ C. The trypsin is neutralized by the feeder medium, which contains 10% FBS. The attached cells that are obtained by trypsinization in this step are the hESC-derived feeders, which have outgrown from the hESC colonies and differentiated into a monolayer of fibroblast-like cells.

7. Remove the supernatant and resuspend the hESC-derived feeders with 12 ml feeder medium and transfer into one gelatin-coated 75-cm2 flask. The number of cells will vary because of culture variability. At this point, you want the most cells possible. Ideally, the hESC-derived feeders are plated at a 1.3 × 104 cells/cm2 density. Therefore, the size of the culture vessel (flask or well) is selected depending on the cell number obtained and the number of hESC-derived irradiated feeder vials to be prepared.

8. Allow the cells to grow to 70% to 80% confluency (∼7 days) before passaging cells (Fig. 1C.5.1C).

Passage hESC-derived feeder cells 9. When the cells are 70% to 80% confluent, aspirate the feeder cell medium. 10. Wash the flask with 5 to 10 ml of CMF-DPBS. 11. Add 3 to 4 ml of trypsin/EDTA to the flask and incubate at 37◦ C for 3 to 5 min. 12. Gently shake the flask to remove the cells, add 10 ml of feeder cell medium, rinse the surface of the flask, and transfer 5 ml to each of two new 75-cm2 flasks. 13. Add 10 ml of feeder cell medium to all three flasks and return to the incubator for further culture. 14. Monitor the culture daily and passage 1:3 when the cultures are 80% to 90% confluent.

Autogenic Human Feeder Cells for hESCs

Inactivate hESC-derived feeder cells Cells can be passaged up to 20 times and inactivated when they reach 90% to 100% confluency.

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To inactivate by γ irradiation 15a. Harvest the feeder layer by aspirating the supernatant and adding 3 to 4 ml trypsin/EDTA, then incubating 3 to 5 min at 37◦ C. After incubation, add an equal volume of feeder cell medium to inactivate the trypsin. 16a. Remove the cells from the flask, transfer them to a 15-ml conical tube and wash by centrifuging at 350 × g, 4◦ C, removing the supernatant, adding 10 ml feeder cell medium, centrifuging again as before, and removing the supernatant. 17a. Irradiate for a total of 40 Gy (4000 rad) using a γ irradiator. 18a. Following irradiation, dilute the feeders to 3 × 105 cells/ml in feeder cell medium. Replate the cells on the appropriate size/number of gelatinized culture dishes to provide feeders to meet experimental goals, and incubate overnight. The inactivated hESC-derived feeder cells are plated on gelatinized dishes in order to provide better support for the long culture periods required for hESC culture. For a 6-well plate, add 1 ml/well of the cell suspension; for a 10-cm dish, add 6 ml of the suspension.

19a. The next day, aspirate the feeder cell medium, rinse culture vessel with CMFDPBS, and replace the culture medium with a volume of hESC medium equal to the previous volume of feeder cell medium.

To inactivate by mitomycin C treatment CAUTION: Mitomycin C is a cytotoxic antitumor agent and must be handled carefully. It works by cross-linking the DNA, which blocks cell division. Handlers should wear latex or nitrile protective gloves and work in a biological safety or fume hood. 15b. Aspirate the feeder cell medium. Add 10 ml of mitomycin C medium per 75 cm2 of culture surface. Incubate 3 hr. Make sure the entire flask is covered with mitomycin C medium so that the inactivation is complete and all cells are exposed for the entire incubation time.

16b. Aspirate mitomycin C solution and inactivate the removed solution with 200 µl bleach. 17b. Wash inactivated feeder layer three times, each time with 10 ml CMF-DPBS. 18b. Harvest the cells with trypsin/EDTA using the technique described in step 15a. Resuspend the cells in 10 ml of feeder cell medium, replate on the appropriate size/number of gelatin-coated 6-well plates, and incubate overnight. 19b. The next day, rinse the culture vessels with 2 ml per well CMF-DPBS (for 6-well plate) and refeed with 5 ml per well of either feeder cell medium or hESC medium. 20. Replate the inactivated feeders on gelatin-coated 175-cm2 flasks and allow them to attach in the incubator for at least 4 hr before culturing with ESCs. The inactivated hESC-derived feeder cells are plated on gelatinized dishes in order to provide better support for the long culture periods required for hESC culture.

Freeze stock hESC-derived feeders 21. At the end of the procedure for preparing inactivated fibroblasts (step 19a or b), aspirate the medium. Cells should be 90% confluent at the time of freezing.

22. Wash the 175-cm2 flask with 10 ml of CMF-DPBS. 23. Add 5 ml of trypsin/EDTA to the flask and incubate at 37◦ C for 3 to 5 min.

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24. Gently shake the flask to lift the cells from the culture surface and neutralize the trypsin by adding 10 ml feeder cell medium. 25. Transfer the entire cell suspension to a 15-ml conical tube and centrifuge tube 5 min at 350 × g, 4◦ C. 26. Remove supernatant and resuspend cells with 1 ml freshly made cold freezing medium. 27. Transfer cell suspension to a 1-ml cryovial. 28. Place vial in an isopropanol freezing container and put container in a −80◦ C freezer overnight. 29. Transfer vial to liquid nitrogen. SUPPORT PROTOCOL 1

PASSAGING hESCs ONTO hESC-DERIVED FEEDERS BY MECHANICAL DISSOCIATION hESCs, unlike mouse ESCs, do not survive well when dissociated to single cells. Therefore, the most reliable method for passaging undifferentiated hESC cultures is manual dissection of the colonies. This method may seem tedious, but it is virtually foolproof and we recommend that novices use this method until they have familiarity with the cells and can easily recognize differentiation in the culture. We also recommend manual passaging for producing cell banks of low-passage hESCs. Enzymatic dissociation methods are provided as an alternative (see Support Protocol 2).

General recommendations a. Passage the cells at a split ratio of ∼1:3 (dividing one dish among three new dishes of the same size) every 5 to 7 days. b. Prepare the feeder layer or extracellular matrix (ECM) substratum the day before passaging. c. There will be considerable variation in the size of colonies in a single dish. Compared with their mouse counterparts, hESCs do not substantially pile up on each other, and their colonies can grow to a large diameter while remaining undifferentiated. Culture conditions affect the flatness of the colonies, but, as an approximation, they are ready to be split when the diameter fills the 10× field when observed under the microscope, as shown in Figure 1C.5.2. d. Examine the culture daily for colony morphology under the phase-contrast or dissecting microscope. e. To be certain that the colonies selected are undifferentiated, it is advisable to dissect the colonies while viewing the dish under a dissecting microscope with illumination from the base. But this is not absolutely necessary, and some prefer to passage the cells without magnification.

Materials hESC cultures hESC medium (see recipe) Stock hESC-derived feeder cells, irradiated or mitomycin C-treated (Basic Protocol) Autogenic Human Feeder Cells for hESCs

Microscope: inverted phase-contrast with 4×, 10×, 20× objectives 15-ml conical centrifuge tubes Pipettor with 20-µl sterile filter tip, or 23-G needle

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Figure 1C.5.2 (A) Phase-contrast image of an undifferentiated hESC (WA09) colony growing in feeder-free culture conditions with MEF conditioned medium. Black arrows point at fibroblast-like cells that spontaneously emerge from the edges of hESC colonies during feeder-free culture. (B) Phase-contrast image of a culture where all of the undifferentiated hESC colonies are completely mechanically dissected away and hESC-derived feeder cells are left behind. (C) Phasecontrast image of a culture of isolated hESC-derived feeders.

1. Evaluate the hESC culture under 4× or 10× phase-contrast optics. The cells can be split among three to six dishes of the same size as the original culture, depending on the density of the original culture. If you wish to put the cells in differentsized dishes, calculate the volume to add based on surface area of each type of dish. The split ratio depends on the number of hESC colonies that are dissociated. If >30 colonies, split 1:6 (i.e., into six new vessels of the original size); if <30 colonies, split 1:3 (i.e., into three new vessels of the original size).

2. Mark (by circling on the bottom of the tissue culture surface) or remove overtly differentiated colonies so as not to disturb these during the dissociation process.

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3. Remove the medium from the dish (or wells) and replace with an equal volume of fresh hESC medium. 4. Dissect the colonies by hand, either under a low-power dissecting microscope (in a horizontal flow hood). A number of different implements can be used to slice up or break up the colonies. Because they are inexpensive and sterile, we recommend either a 20-µl pipettor with a sterile filter tip attached, or a sterile 23-G needle.

5. Cut the colony into strips, and then into squares, so that each piece of the colony has a few hundred cells. 6. Break up each colony by moving the tip around and across each colony in a spiral motion. 7. After all of the colonies are dissected, use a 5-ml pipet to transfer the culture medium containing the dissected colonies to a 15-ml conical tube. Rinse the plate with 5 ml hESC medium and add this to the same 15-ml tube. 8. Bring up the final volume in the tube to 8 to 10 ml with hESC medium. 9. Gently triturate the cell clumps into smaller clumps of ∼30 cells using a sterile 10-ml pipet and divide the suspension into the prepared culture dishes containing stock irradiated or mitomycin C–treated hESC-derived feeder layers (see Basic Protocol) in the appropriate amount of hESC medium. SUPPORT PROTOCOL 2

PASSAGING hESCs ONTO hESC-DERIVED FEEDERS BY ENZYMATIC DISSOCIATION As an alternative to manual passaging (see Support Protocol 1), passaging may be performed following enzymatic dissociation of the cells. The advantage of this procedure relative to manual passaging is that it need not be performed under a dissecting microscope. Enzymatic dissociation methods vary widely, and the exact conditions need to be developed for each laboratory. Most importantly, cultures that have been maintained by manual passaging cannot be passaged by enzymatic dissociation unless exceptional care is taken to adapt and monitor the cells. The type of enzyme used for dissociation is critical. For example, passaging with trypsin appears to put more selective pressure on the cultures than other methods, resulting in a higher incidence of drift of hESC lines toward aneuploidy. But some hESC lines have been derived using trypsin from the outset. These lines can be routinely passaged using whatever enzymatic technique is provided by the supplier. Microbial collagenase is preferred by many laboratories, perhaps because of the way in which it is used. Collagenase is used by many investigators to loosen the hESC colonies from the dishes, not to dissociate them to single cells, and the cell clumps have to be further dissociated by trituration. Keep in mind that enzymes are not highly purified recombinant products, and they may contain animal products. Trypsin is prepared from porcine tissue, and collagenase is a crude microbial product.

Materials

Autogenic Human Feeder Cells for hESCs

1C.5.8 Supplement 4

hESC cultures Calcium and magnesium-free Dulbecco’s phosphate-buffered saline (CMF-DPBS; e.g., Sigma or Invitrogen) 200 U/ml collagenase IV (see recipe) hESC medium (see recipe) Current Protocols in Stem Cell Biology

Stock hESC-derived feeder cells, irradiated or mitomycin C-treated (Basic Protocol) 15-ml conical tubes 1. Remove the hESC culture medium. 2. Rinse culture with 10-ml CMF-DPBS. 3. Add sufficient 200 U/ml of collagenase IV to cover the cells and incubate 5 to 10 min at 37◦ C, until the edges of the colonies start to curl up. Observe the culture under the microscope. 4. Remove the collagenase and replace with 2 ml of hESC medium (if using a 6-well plate or 35-mm dish). Adjust the volume of medium used depending on the size of the culture dish/well/flask.

5. Using a 5-ml pipet, gently dislodge the colonies from the plate and place them in a 15-ml conical tube. 6. Gently triturate the cell clumps into smaller clumps of ∼30 cells using a sterile 10-ml pipet and divide the suspension into the prepared culture dishes containing stock irradiated or mitomycin C–treated hESC-derived feeder layers (see Basic Protocol) in the appropriate amount of hESC medium. The cells can be split into three to six dishes of the same size as the original culture, depending on density of the original culture. If you wish to put the cells in different sized dishes, calculate the dilution based on the surface area of each type of dish.

GROWTH OF hESCs UNDER FEEDER-FREE MEF-CM CULTURE CONDITIONS

SUPPORT PROTOCOL 3

This protocol describes the transfer of hESC cultures growing on mouse embryonic fibroblast (MEF) feeders to a feeder-free hESC culture system that consists of a MEFconditioned hESC medium (MEF-CM) and a BD Matrigel substratum. A similar strategy, using human feeder cell conditioned medium, may be adopted for hESCs grown on human feeder cell layers. MEF feeders are cultured and inactivated by γ irradiation or mitomycin C. There are several commercial sources for MEFs from various mouse strains containing various selectable drug-resistant markers [ATCC (http://www.atcc.org); Chemicon International (http://www.chemicon.com); Primogenix, Inc. (http://www. primogenix.com); Stem Cell Technologies (http://www.stemcell.com); GlobalStem, Inc. (http://www.globalstem.com)]. Depending on the level of use, this can be a convenient and economical alternative to the de novo preparation of MEF. For culturing in MEF-CM, the hESCs must be plated on Matrigel. BD Matrigel is derived from the Engelbreth-Hom-Swarm mouse tumor cell line. It is very rich in extracellular matrix components and comprises ∼60% laminin, 30% collagen IV, 8% entactin, heparin sulfate proteoglycan, and low levels of growth factors. It is a liquid at 4◦ C, but polymerizes quickly at room temperature. Growth of hESC cultures in MEF-CM to allow spontaneous differentiation of hESCs into fibroblast-like cells requires 5 to 7 days.

Materials MEFs (see above), inactivated by γ irradiation or mitomycin C treatment Feeder medium (see recipe) hESC medium (see recipe) containing 4 ng/ml human FGF2

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10 µg/ml human FGF2 (see recipe) Matrigel (BD Biosciences) KnockOUT DMEM medium (Invitrogen) 10-cm culture dishes (∼55 cm2 area) 0.2-µm low-protein-binding filters 1-, 10-, 15-, 25-, and 50-ml tubes 6-well tissue culture plates, prechilled Inverted phase-contrast microscope Low-power dissecting microscope Pipettor with 20-µl sterile filter tip, or 23-G needle Prepare MEF-conditioned medium (MEF-CM) 1. Plate inactivated MEFs at 1 × 106 cells per 10-cm culture dish in feeder medium and allow them to attach overnight in the incubator. Feeder medium contains fetal bovine serum (FBS), and MEFs will not attach well to the tissue culture flask in hESC medium without FBS.

2. The next day, remove the feeder cell medium, rinse the cell layer with hESC medium containing 4 ng/ml human FGF2, then replace feeder medium with 27.5 ml hESC medium (0.5 ml/cm2 ) containing 4 ng/ml human FGF2. Use ∼ 27.5 ml of hESC medium for a 10-cm tissue culture dish (surface area, ∼55 cm2), 38 ml of hESC medium for a 75-cm2 flask, 75 ml for a 150-cm2 flask, or 112 ml for a 225-cm2 flask.

3. Allow MEFs to condition the hESC medium for 24 hr. 4. Collect conditioned hESC medium (MEF-CM) from feeder cell flask daily for up to 7 days. Harvest MEF-CM at about the same time each day in an effort to minimize the variability of the medium.

5. Replace the medium in the flasks with fresh hESC medium. 6. Filter MEF-CM through a 0.2-µm low-protein-binding filter and divide into 10-ml, 25-ml, and 50-ml aliquots in labeled and dated sterile tubes. Store up to 6 months at −20◦ C. Prior to using MEF-CM to feed hESC cultures, supplement with an additional 8 ng/ml of human FGF2.

Prepare Matrigel stock solution 7. The evening prior to preparation of the stock solution, thaw Matrigel bottle slowly, overnight at 4◦ C. Slow overnight thawing at 4◦ C is necessary to prevent the Matrigel from polymerizing. Keep thawed Matrigel on ice until use. When preparing gel, use precooled pipets, plates, and tubes. BD Matrigel Matrix will gel rapidly at 22◦ to 33◦ C.

8. Add 10 ml of cold KnockOUT DMEM to the bottle containing 10 ml of Matrigel. 9. Keeping the mixture on ice, mix well by pipetting up and down with a 10-ml pipet. 10. Aliquot 1 to 2 ml into pre-chilled 1-ml tubes (on ice) and store aliquots at −20◦ C.

Prepare Matrigel-coated plates 11. Slowly thaw a frozen Matrigel aliquot at 4◦ C on ice. Autogenic Human Feeder Cells for hESCs

12. Dilute Matrigel aliquots 1:15 in cold KnockOUT DMEM medium (1:30 final). 13. Add 1 ml of diluted Matrigel/well of a 6-well plate.

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14. Incubate the plates for at least 1 hr at room temperature or overnight at 4◦ C. If stored at 4◦ C, coated plates can be used for up to 1 week after coating.

Passage hESCs onto Matrigel-coated plates 15. Evaluate the hESC culture to be passaged under 4× or 10× phase contrast microscopy. Culture is ready to be split when the diameter of a hESC colony fills the 10× field when observed under the microscope, as shown in Figure 1C.5.1. Usually, hESC cultures are ready to be passaged by day 5 to 7 of culture. The cells can be split between two dishes of the same size as the original culture, depending on the density of the original culture. If you wish to put the cells in different sized dishes, calculate the volume to add based on the surface area of each type of dish.

16. Mark (or remove) overtly differentiated colonies so as not to disturb these during the dissociation process. 17. Remove the medium from the dish and replace with an equal volume of MEF-CM (supplemented with an additional 8 ng/ml human FGF2; see step 6). 18. Dissect the colonies by hand, under a low-power dissecting microscope (in a horizontal flow hood). Several implements can be used to slice up or break up the colonies. Because they are inexpensive and sterile, we recommend either a 20-µl pipettor with a sterile filter tip attached, or a sterile 23-G needle.

19. Cut the colony into strips, and then into squares so that each piece of the colony has a few hundred cells. 20. Break up each colony by moving the tip around and across each colony in a spiral motion. 21. After all of the colonies are dissected, use a 5-ml pipet to transfer the culture medium containing the dissected colonies to a 15-ml conical tube. Rinse the plate with MEFCM (supplemented with an additional 8 ng/ml human FGF2; see step 6) and add this to the same 15-ml tube. 22. Bring up the final volume in the tube to 8 to 10 ml with MEF-CM (supplemented with an additional 8 ng/ml human FGF2; see step 6). 23. Just before transferring the dissected hESC colonies, remove the Matrigel-containing solution from the culture plates. 24. Gently triturate the cell clumps using a sterile 10-ml pipet, and divide the suspension into the prepared culture Matrigel coated dishes, using a split ratio of 1:2 (i.e., into two dishes).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture-grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Collagenase IV, 200 U/ml Dissolve 20,000 U of microbial collagenase IV (Invitrogen) in 100 ml of KnockOUT DMEM (Invitrogen). Filter using a 250-ml 0.2-µm, low-protein-binding filter. Aliquot and store at −20◦ C. Alternatively, make 1 mg/ml solution in KnockOUT DMEM, filter, aliquot, and store at −20◦ C. This solution is used in the protocol at the same volume as that described above.

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FGF2, 10 µg/ml Dissolve 10 µg human FGF2 (Millipore) in 1 ml of CMF-DPBS containing 0.2% (w/v) BSA. Divide into 50- to 100-µl aliquots. Store frozen at −20◦ C or −80◦ C for long-term storage. Thawed aliquots are stable for up to 2 weeks at 4◦ C.

Feeder medium 435 ml high-glucose DMEM+GlutaMax (Invitrogen) 50 ml fetal bovine serum (FBS; 10% final) 5 ml 100× nonessential amino acids (Invitrogen; 1× final) 5 ml 100× penicillin-streptomycin (optional; Invitrogen; 1× final) Store up to 3 months at −20◦ C hESC medium 400 ml high-glucose DMEM/F12/Glutamax (Invitrogen) 100 ml KnockOUT serum replacement (Invitrogen; 20% final) 5 ml 100× nonessential amino acids (Invitrogen; 1× final) 910 µl 55 mM 2-mercaptoethanol (0.1 mM final) 5 ml 100× penicillin-streptomycin (optional; Invitrogen; 1× final) 200 µl 10 µg/ml human FGF2 (see recipe; 4 ng/ml final) Mix all ingredients, except FGF2, in a 500-ml bottle in a tissue culture hood using aseptic technique, and then filter using a 0.2-µm, low-protein-binding filter unit into a sterile 500-ml bottle. Add FGF2 to medium. Store at 4◦ C. Discard unused medium after 2 weeks.

Mitomycin C medium Remove plastic covering from lid of one vial of mitomycin C (Sigma, cat. no. M-0503). Wipe top with 70% ethanol. Using a 22-G needle attached to a 3-ml syringe, draw up 1 ml CMF-DPBS (Invitrogen or Sigma). Transfer the CMF-DPBS to the vial of mitomycin C powder and swirl vial to dissolve. When dissolved, draw up the solution and carefully detach the syringe, leaving the needle in the vial. Transfer to a 15-ml tube. Wrap tube with foil and store at 4◦ C.

COMMENTARY Background Information

Autogenic Human Feeder Cells for hESCs

Most hESC lines have been derived and maintained using medium containing fetal bovine serum (FBS) or bovine-serum-derived products (such as Invitrogen’s KnockOUT serum replacement, KSR) and coculture with MEFs or mouse-derived Matrigel extracellular matrix. However, concerns about problems with xenografts have motivated development of alternative culture systems that reduce the requirement for animal-derived products. Several groups have begun to use human cells as feeder layers (Table 1C.5.1). Recently, we and others have found that fibroblast-like cells derived from hESCs can be used as an autogeneic feeder system that efficiently supports the growth and maintenance of pluripotency of both autogeneic and allo-

geneic undifferentiated hESCs (Table 1C.5.1). There are several potential advantages to using hESC-derived feeder cells: 1. Derivation and culture of hESC-derived feeders is not time consuming. 2. Expensive tissue biopsy and unnecessary sacrifice of animals are not required. 3. They afford an inexpensive method to eliminate transfer of pathogens from feeders. 4. They provide a good system to study stem cell–feeder “niche” interactions. The karyotype of hESC-derived feeder cells is normal. Microarray analysis indicates that they are differentiated (no detectable OCT4, Sox2, or Nanog expression). High expression of IGF and TGF-B growth factors correlates with good support capacity as feeder cells. Growth of hESC-derived feeders begins to slow down beyond passage 20.

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Table 1C.5.1 Examples of Feeder Cells Derived from Human Tissue Sources that Support Pluripotent Growth of hESCsa

Human feeder cell type

hESC tested

Reference

Comments

hFS (fetal skin fibroblast)

HES3

Richards et al. (2002, 2003)

First to report the use of humanized culture system

ATCC: D551/CCL-10 hFM (fetal muscle fibroblast)

HES4

Human feeders and human serum used for culture

hAFT (adult fallopian tube epithelial cells)

All three of these human fibroblast sources support hESC growth Derivation of new line on hFM cells: human feeders have greater proliferation than MEFs, but limited availability

Human foreskin fibroblast

TE03

Amit et al. (2003)

Longer proliferation potential in culture than MEFs, >42 passages

ATCC: Hs27 (CRL1634)

TE06

ATCC: Hs68 (CRL1635)

WA09

Fetal foreskin fibroblast

HES2

ATCC: BJ (CRL 2522)

HES3

All three feeder lines supported long-term growth, in KnockOUT DMEM, 15% KSR, 4 ng/ml FGF2

ATCC: Hs27 (CRL-1634)

HES4

None supported growth in DMEM, 20% FBS

KnockOUT DMEM, 15% KSR, 4 ng/ml FGF2 Choo et al. (2004)

Evaluated growth of hES lines in FBS and KSR on each of the commercially available hFF lines

ATCC: Hs68 (CRL-1635) Adult bone marrow stromal cells

WA01

Cheng et al. (2003) Supports undifferentiated growth, but stromal cells need to be prepared frequently as they senesce with increasing time in culture as do MEFs

hUECs (uterine endometrial cells)

Miz hES1

Lee et al. (2004)

hBPCs (breast parenchymal cells)

Noted hESC colonies are flatter and thinner on UECs and EFs than on MEFs or hBPCs DMEM/F12, 20% KSR, 4 ng/ml FGF2

hEFs (embryonic fibroblast) hUECs (uterine endometrial cells)

Miz Hes9, 14, 15

Lee et al. (2005)

hUEC “endo-1” line was used to derive new hESC DMEM/F12, 20% KSR, 4 ng/ml FGF2

PLFb (human placental fibroblast)

UC01

Genbacev et al. (2005)

Found human placental fibroblast to be equivalent to MEFS (CF-1)

UC06 MEFs (CF-1)

WA01

Derived a new line on this feeder: UCSF-1

WA09

KnockOUT DMEM, 20% KSR, 4 ng/ml FGF2 continued Embryonic and Extraembryonic Stem Cells

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Table 1C.5.1 Examples of Feeder Cells Derived from Human Tissue Sources that Support Pluripotent Growth of hESCsa , continued

Human feeder cell type

hESC tested

Reference

Comments

HEF1-hTERT hESC-derived (fibroblast-like feeder cells, immortalized by infection with retrovirus expressing hTERT)

WA01

Xu et al. (2004)

First to report genetically modified hESC-derived feeders

WA07 WA09 hESC-df (hESC-derived fibroblast-like feeder cells)

WA01

Stojkovic et al. (2005)

hESC-df support undifferentiated growth of hESCs when cocultured directly, when used to condition hESC medium, and when cultured on Matrigel. Was used to derive a new hESC line.

Diff-Miz-hES6 (hESC-derived fibroblast-like feeder cells)

Miz-hES1

Yoo et al. (2005)

Diff-Miz-hES6 supported growth of hESCs

Stewart et al. (2006)

hDFs supported the growth of FACS-sorted hESCs

Miz-hES4 UC06 hdf (autologously derived hESC-derived fibroblast-like cells)

WA01 WA09

a Abbreviations: ATCC, American Type Culture Collection; DMEM, Dulbecco’s Modified Eagle Medium; EF, embryonic fibroblast; FACS, fluo-

rescence activated cell sorting; FBS, fetal bovine serum; FGF2, fibroblast growth factor 2; hBPC, human breast perenchymal cell; hESC, human embryonic stem cell; hFF, human foreskin fibroblast; KSR, KnockOUT serum replacement (Invitrogen); MEF, mouse embryonic fibroblast.

Critical Parameters and Troubleshooting Strain of mouse used to make MEF-CM MEFs from several mouse strains have been found to support hESC culture. However, the strains currently favored are isolated from the CF-1 or 129 strains. Other strains that have been used to support hESC growth in coculture are FVB/N, B6/129 hybrids, and C57BL/6. The most critical variable seems to be the quality of the MEFs, which should be used between passage 3 and passage 6 and before the culture consists of many large multinucleated fibroblasts. CF-1 MEFs require a higher dose of radiation to inactivate them (60 to 80 Gy) than 129 or B6 MEFs (30 to 40 Gy).

Autogenic Human Feeder Cells for hESCs

Adapting hESC cultures to grow on Matrigel in MEF-CM Transferring cells from fibroblast coculture to the Matrigel/MEF-CM system may require a couple of passages to allow the cells to adapt to culture without feeders. Passage to low-density MEF feeders on BD Matrigel and MEF-CM for the first couple of passages may ease the transition to feeder-free culture.

The hESCs remain karyotypically normal when grown on hESC-derived feeders. We have found no differences in their ability to grow or differentiate. Microarray analysis has also indicated no difference in gene expression between hESCs growing on MEFs and those on hESC-derived feeders. hESC can be carried on Matrigel in MEF-CM for at least 20 passages. Matrigel The source of this ECM mixture is the Engelbreth-Holm-Swarm mouse tumor. Its major components are laminin, collagen IV, heparin sulfate proteoglycans, and entactin. At room temperature, BD Matrigel matrix polymerizes to produce a biologically active matrix material resembling the mammalian cellular basement membrane. While quality control measures are used to minimize the variability between production lots, lot-to-lot variations are inherent in any cell-derived product. Culture results may vary depending on the production lot, the concentration that is plated, the length of time the plates are incubated with Matrigel, and how the plates are stored prior

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to use. An alternative to preparing Matrigelcoated plates in the laboratory is to purchase precoated plates, which have been coated with an optimized concentration and prepared and tested for culture with hESCs. Contamination of cultures With good culture technique, bacterial contamination should not be a problem. We recommend that antibiotics be used while new investigators are being trained in the techniques. Antibiotics such as penicillin and streptomycin do not have any effect on mycoplasma. Mycoplasma is a serious problem in laboratories that culture multiple cell lines or that have inadequately trained personnel.

Anticipated Results We found that hESC-derived feeder cells will maintain hESC lines in an undifferentiated, karyotypically intact state. These autologous hESC-derived fibroblast-like feeders can be easily accessed by researchers, and aid in the development of a xenofree hESC culture system, thus promoting the safety of cell replacement therapy.

Time Considerations Growth of hESC cultures in MEF-CM to allow spontaneous differentiation of hESCs into fibroblast-like cells requires 5 to 7 days. Isolation, expansion, and inactivation of hES-derived feeders requires 7 days. Growth of hESCs on hESC-derived feeder cells requires 5 to 7 days per passage.

Literature Cited Amit, M., Margulets, V., Segev, H., Shariki, K., Laevsky, I., Coleman, R., and Itskovitz-Eldor, J. 2003. Human feeder layers for human embryonic stem cells. Biol. Reprod. 68:2150-2156. Cheng, L., Hammond, H., Ye, Z., Zhan, X., and Dravid, G. 2003. Human adult marrow cells support prolonged expansion of human embryonic stem cells in culture. Stem Cells 21:131-142.

Genbacev, O., Krtolica, A., Zdravkovic, T., Brunette, E., Powell, S., Nath, A., Caceres, E., McMaster, M., McDonagh, S., Li, Y., Mandalam, R., Liebowski, J., and Fisher, S.J. 2005. Serum-free derivation of human embryonic stem cell lines on human placental fibroblast feeders. Fertil. Steril. 83:1517-1529. Lee, J.B., Song, J.M., Lee, J.E., Park, J.H., Kim, S.J., Kang, S.M., Kwon, J.N., Kim, M.K., Roh, S.I., and Yoon, H.S. 2004. Available human feeder cells for the maintenance of human embryonic stem cells. Reproduction 128:727-735. Lee, J.B., Lee, J.E., Park, J.H., Kim, S.J., Kim, M.K., Roh, S.I., and Yoon, H.S. 2005. Establishment and maintenance of human embryonic stem cell lines on human feeder cells derived from uterine endometrium under serumfree condition. Biol. Reprod. 72:42-49. Richards, M., Fong, C.Y., Chan, W.K., Wong, P.C., and Bongso, A. 2002. Human feeders support prolonged undifferentiated growth of human inner cell masses and embryonic stem cells. Nat. Biotechnol. 20:933-936. Richards, M., Tan, S., Fong, C.Y., Biswas, A., Chan, W.K., and Bongso, A. 2003. Comparative evaluation of various human feeders for prolonged undifferentiated growth of human embryonic stem cells. Stem Cells 21:546-556. Stewart, M.H., Bosse, M., Chadwick, K., Menendez, P., Bendall, S.C., and Bhatia, M. 2006. Clonal isolation of hESCs reveals heterogeneity within the pluripotent stem cell compartment. Nat. Methods 3:807-815. Stojkovic, P., Lako, M., Przyborski, S., Stewart, R., Armstrong, L., Evans, J., Zhang, X., and Stojkovic, M. 2005. Human-serum matrix supports undifferentiated growth of human embryonic stem cells. Stem Cells 23:895-902. Xu, C., Jiang, J., Sottile, V., McWhir, J., Lebkowski, J., and Carpenter, M.K. 2004. Immortalized fibroblast-like cells derived from human embryonic stem cells support undifferentiated cell growth. Stem Cells 22:972-980. Yoo, S.J., Yoon, B.S., Kim, J.M., Song, J.M., Roh, S., You, S., and Yoon, H.S. 2005. Efficient culture system for human embryonic stem cells using autologous human embryonic stem cellderived feeder cells. Exp. Mol. Med. 37:399407.

Choo, A.B., Padmanabhan, J., Chin, A.C., and Oh, S.K. 2004. Expansion of pluripotent human embryonic stem cells on human feeders. Biotechnol. Bioeng. 88:321-331.

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1C.5.15 Current Protocols in Stem Cell Biology

Supplement 4

Isolation of Human Placental Fibroblasts 1

2

UNIT 1C.6

2

Dusko Ilic, Mirhan Kapidzic, and Olga Genbacev 1 2

StemLifeLine, San Carlos, California University of California, San Francisco, California

ABSTRACT The first human embryonic stem cell lines (hESCs) were derived using mouse embryonic fibroblasts as feeder cells. In attempts to replace mouse embryonic fibroblasts with feeders of human origin, irradiated human placental fibroblasts were successfully used as feeder cells for the derivation and propagation of hESCs. Here we describe a protocol for the isolation and expansion of fibroblasts from placental villous stroma. We include a description of placental architecture to provide the background for a stepwise tissue digestion that leads to the isolation of villous stroma. Villous stroma from the first trimester tissue is different from term placenta and contains mesenchymal, fibroblastlike cells, only a few blood vessels, and a network of matrix fibers. The fibroblasts isolated from a single placenta of 6- to 8-weeks gestation proliferate rapidly and retain the ability to support hESC growth between passage doubling (PD) 8 and PD 12. Curr. C 2008 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 5:1C.6.1-1C.6.17.
Keywords: placenta r fibroblasts r human feeders r human embryonic stem cells

INTRODUCTION The first human embryonic stem cell lines (hESCs) were derived using mouse embryonic fibroblasts as feeder cells. In attempts to replace mouse embryonic fibroblasts with feeders of human origin, irradiated human placental fibroblasts were successfully used as feeder cells for the derivation and propagation of hESCs. This unit describes protocols for collecting placentas and isolating placental fibroblasts (Basic Protocol 1), expanding and freezing placental fibroblasts (Basic Protocol 2), dressing appropriately for these procedures (Support Protocol 1), and counting the cells using a cell counter (Support Protocol 2). NOTE: All studies with human subjects must be approved by the Institutional Review Board (IRB), which must adhere to the Office for the Protection from Research Risk (OPRR) guidelines or other applicable governmental regulations for using human subjects. All material must be obtained with informed consent of the donor. NOTE: All procedures should be performed under sterile conditions. All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly.

COLLECTION OF PLACENTAL TISSUE AND ISOLATION OF PLACENTAL FIBROBLASTS Human placentas (6 to 8 weeks of gestation) are obtained individually with informed consent and using a protocol approved by the Institutional Review Board or equivalent committee. Tissues from patients positive for human immunodeficiency virus, hepatitis, or sexually transmitted diseases are not collected. Isolation of human placental fibroblasts is a stepwise process during which the villi are subjected to five enzymatic solutions.

Current Protocols in Stem Cell Biology 1C.6.1-1C.6.17 Published online June 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c06s5 C 2008 John Wiley & Sons, Inc. Copyright


BASIC PROTOCOL 1

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1C.6.1 Supplement 5

Figure 1C.6.1 Step-wise tissue disaggregation. (A) Dissected placental villi. (B) After the first collagenase digestion digested syncytiotrophoblast, blood cells, and other debris will remain floating in a collagenase supernatant. (C) After the first trypsin digestion, detached cells, mostly cytotrophoblasts, and some fibroblasts will remain floating in a trypsin supernatant. (D) Fibroblasts are ∼20% of detached cells that remain floating in a trypsin supernatant after second trypsin digestion. Majority of detached cells that remain floating in a supernatant after the third trypsin (E) and after the second collagenase digestion (F) are fibroblasts. Drawing: Joseph Hill, Hill-SciArt.

Each solution disaggregates a particular cell layer of the tissue, as shown in Figure 1C.6.1 and described in the protocol.

Materials Placental tissue Phosphate-buffered saline (PBS), Ca2+ -, Mg2+ -free (CMF-PBS; Invitrogen, cat. no. 14190-144) Cytowash medium (see recipe) Ice bucket or Styrofoam box filled with ice 70% ethanol (Fisher Scientific, cat. no. 254670–32) Collagenase solution (see recipe) Trypsin solution (see recipe) Fetal bovine serum (Hyclone, cat. no. SH30071.03) Fibroblast culture medium (see recipe)

Isolation of Human Placental Fibroblast Feeder Cells

Full height sieves, stainless steel wire and frame, Tyler (VWR Lab Shop, cat. no. 57324-181), 1-mm mesh opening 15-cm petri dish White Light Transilluminator (Fisher Scientific; Fisher Biotech, cat. no. FB-WLT-1417) Sterile surgical instruments: Forceps (Fisher, cat. no. 10275)

1C.6.2 Supplement 5

Current Protocols in Stem Cell Biology

Dumont tweezers, high precision, 120 mm, stainless steel (Ted Pella, cat. no. 5617) General laboratory scissors, curved with sharp points, 110 mm, stainless steel (Ted Pella, cat. no. 1332) Vannas-type Micro Scissors, straight, 80 mm, stainless steel (Ted Pella, cat. no. 1346) 50-ml polypropylene conical tube (BD Falcon, cat. no. 352098) 37◦ C shaking water bath Centrifuge 145 × 20–mm cell culture dish (USA Scientific, cat. no. 5663-9160) 100-ml heat-resistant glass bottle (Corning, cat. no. X1395), sterile Medical sterile gauze sponges, 2 × 2–in. (Kendall Curity Gauze Sponge, cat. no. 1806) 100-µm cell strainer (Fisher Scientific, cat. no. 08-771-19), sterile 25-cm2 flasks Additional reagents and equipment for dressing for collection and isolation of placental fibroblasts (Support Protocol 1) and counting cells (Support Protocol 2) NOTE: One set of sterile instruments and labware listed above is required for each placenta processed. Catalog numbers for the surgical instruments and labware are given just as examples—instruments and labware of the equivalent quality can be purchased from other manufacturers without altering the outcome of the procedure.

Collect placental tissue 1. Don protective clothing following the procedure described in Support Protocol 1. 2. Place the tissue into sterile sieve and wash the tissue with 50 ml ice-cold CMF-PBS. 3. Repeat the procedure as many times as needed to get rid of the blood. 4. Place tissue in 15-cm petri dish on transilluminator and remove coagulated blood using forceps and scissors. 5. Transfer the tissue into a 50-ml conical tube and add cytowash medium to cover the tissue. 6. Place the tube on ice. 7. Repeat steps 1 to 5 with another sample using a new sterile set of the surgical instruments and sterile dishes. IMPORTANT NOTE: To avoid cross-contamination do not mix tissue samples or use the same instruments or dishes for two different placentas. Collect as many samples as needed or as many as are available that day. The best practice is to start with two placentas. An experienced person can handle up to four placentas at the same time without jeopardizing the outcome of the prep.

8. Transport samples on ice to the laboratory. The actual time period within which the samples are collected depends also on the distance from the site of collection to the laboratory. Collection and transport should not take longer than 4 to 6 hr from the beginning of collecting the first sample to the beginning of the isolation procedure.

Prepare laboratory equipment 9. Switch shaking water bath on and set temperature to 37◦ C. To keep water in shaking water bath clean add 2 to 3 drops per liter of Bath Clear (Fisher Scientific, cat. no. 13-641-337) once or twice a week.

10. Switch centrifuge on and set temperature to 4◦ C.

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11. Wipe an unplugged white-light transilluminator with a cloth dampened with 70% ethanol and then set it up in a biosafety cabinet. 12. Don protective clothing following the procedure described in Support Protocol 1. 13. Keep samples in a biosafety cabinet on ice. To avoid adventitious contamination of ongoing cell culture it is advisable to have one biosafety cabinet dedicated only for preparation of primary cells directly from tissue samples, whereas all other cell culture work in the laboratory should be done in a separate biosafety cabinet.

14. Place large cell culture dish (145 × 20–mm) on transilluminator, fill up with 20 to 30 ml of cytowash medium. Dissect placental villi 15. Place placenta sample in the culture dish. Separate membrane from placental villi using sterile surgical instruments. 16. Cut villi to 3- to 4-mm pieces and collect them into preweighed sterile 50-ml tube. Figures 1C.6.2 and 1C.6.3 (lower and higher magnification, respectively) show maternal (A) and fetal (B) sides of the placenta before separation of the membranes. Dissected villi are shown in Figure 1C.6.4.

17. Fill the tube with cytowash medium up to 50 ml. 18. Centrifuge the tube 5 min at 700 × g, 4◦ C. 19. Aspirate supernatant and weigh the tube with the tissue. To determine the weight of the tissue, subtract the weight of the empty tube from the total weight. One placenta of 6- to 8-weeks gestational age yields 1 to 3 g of the villous tissue. If the weight is <1 g, stop here and dispose of the sample in medical waste according to the institutional protocol.

Perform first collagenase digestion 20. Add 7 ml collagenase solution per gram of tissue, and transfer suspended tissue into a sterile 100-ml glass bottle. 21. Place the bottle into shaking water bath at 37◦ C. Shake at 175 rpm for 6 min. Assure that the bottle is anchored safely so that it will not tip over.

22. Remove the bottle from the water bath. Spray the bottle with 70% ethanol and place on ice in a biosafety cabinet. Leave the bottle for 2 to 3 min on ice. Under the influence of gravity undigested tissue will fall to the bottom of the bottle, whereas digested syncytiotrophoblast, blood cells, and other debris will remain floating in a collagenase supernatant.

23. Aspirate and discard supernatant. Perform first trypsin digestion 24. Resuspend tissue in 7 ml trypsin solution per gram of tissue, and place the bottle into shaking water bath at 37◦ C. Shake at 175 rpm for 10 min. 25. Remove the bottle from the water bath. Spray the bottle with 70% ethanol and place on ice in biosafety cabinet. 26. Leave the bottle for 2 to 3 min on ice. Isolation of Human Placental Fibroblast Feeder Cells

Under the influence of gravity undigested tissue will fall to the bottom of the tube, whereas detached cells, mostly cytotrophoblast, and some fibroblasts will remain floating in a trypsin supernatant.

27. Aspirate and discard supernatant.

1C.6.4 Supplement 5

Current Protocols in Stem Cell Biology

Figure 1C.6.2 (A) Maternal side of the specimen with chorionic villi (CV) covering the surface. (B) Fetal side of the specimen with chorionic villi (CV) and fetal membranes (FM).

Figure 1C.6.3 (A) Maternal side of the specimen with chorionic villi (CV) covering the surface. (B) Fetal side of the specimen with chorionic villi (CV) and fetal membranes (FM).

Figure 1C.6.4 Placental chorionic villi (CV) separated from membranes, dissected, minced, and rinsed in culture media. BV, blood vessels in villus core. BC, remaining blood clots.

Embryonic and Extraembryonic Stem Cells

1C.6.5 Current Protocols in Stem Cell Biology

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Perform second trypsin digestion 28. Resuspend tissue remaining in the bottle in 7 ml trypsin solution per gram of initial tissue weight as measured in step 19, and place the bottle into shaking water bath at 37◦ C. Shake at 175 rpm for 30 min. 29. Remove the bottle from the water bath. Spray the bottle with 70% ethanol and place on ice in biosafety cabinet. 30. Leave the bottle for 2 to 3 min on ice. Under the influence of gravity undigested tissue will fall to the bottom of the tube, whereas detached cells, mostly fibroblasts, will remain floating in a trypsin supernatant. Fibroblasts are ∼20% of detached cells that remain floating in a trypsin supernatant after second trypsin digestion.

Collect fibroblasts 31. In a clean sterile 50-ml tube add 4 ml fetal bovine serum. 32. Cover the top of the tube with one piece of sterile gauze sponge and pass supernatant from the bottle through the gauze filter. 33. Fill up the tube with cytowash medium up to 50 ml and mix content. 34. Centrifuge the tube 8 min at 700 × g, 4◦ C. 35. Aspirate supernatant and resuspend cell pellet in 10 ml fibroblast culture medium.

Perform third trypsin digestion 36. Resuspend tissue remaining in the bottle in 4 ml trypsin solution per gram of initial tissue weight as measured in step 19, and place the bottle into shaking water bath at 37◦ C. Shake at 175 rpm for 30 min. 37. Repeat steps 29 through 35. The majority of detached cells that remain floating in a trypsin supernatant after the third trypsin digestion are fibroblasts.

Perform second collagenase digestion 38. Resuspend tissue remaining in the bottle in 4 ml collagenase solution per gram of initial tissue weight as measured in step 19, and place the bottle into shaking water bath at 37◦ C. Shake at 175 rpm for 3 min. 39. Repeat steps 29 through 35. The majority of detached cells that remain floating in the supernatant after second collagenase digestion are fibroblasts.

40. Pool together cell suspensions remaining after the second trypsin digestion, the third trypsin digestion, and the second collagenase digestion (total 30 ml) and, to exclude large clumps, pass through a sterile 100-µm cell strainer into 50-ml tube. 41. Add fibroblast medium up to 50 ml. Resuspend well. 42. Centrifuge the tube 8 min at 700 × g, 4◦ C. 43. Aspirate supernatant and resuspend pellet in 5 ml fibroblast culture medium. 44. Count cells (see Supporting Protocol 2) and record the total number of cells on flasks. Isolation of Human Placental Fibroblast Feeder Cells

This number is the starting number or passage doubling (PD) 1.

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Current Protocols in Stem Cell Biology

45. If there are <400,000 cells, plate all cells into a 25-cm2 flask. If there are >400,000 cells plate cells in 250,000 to 400,000 cells per flask. Incubate the flask(s) in cell culture incubator at 37◦ C and 5% CO2 . Normal yield is ∼400,000 to 500,000 cells per placenta. If the yield is <250,000 cells, do not proceed further. Dispose of the cells in medical waste according to the institutional protocol.

DRESSING FOR COLLECTION AND ISOLATION OF PLACENTAL FIBROBLASTS

SUPPORT PROTOCOL 1

Protective clothing is important to protect the investigator from pathogens that might be present in the raw material of human origin. The rules regarding safety and gowning may differ according to the institution. All the samples should be treated as if they are infectious and appropriate protection should be used.

Materials Antibacterial soap Sterile surgical gown (VWR, cat. no. 10845–016) Shoe covers (Fisher Scientific, cat. no. 19–098–921) Hair cover (Fisher Scientific, cat. no. 17–981–43A) Protective glasses (Fisher Scientific, cat. no. 19–130–2088) Surgical face-mask (Fisher Scientific, cat. no. 18–096B) Sterile surgical gloves (Fisher Scientific, cat. no. 18–999–2668) NOTE: Catalog numbers for the protective clothing are given just as examples— protective clothing of the equivalent quality can be purchased from other manufacturers without altering the outcome of the procedure.

Gown properly 1. Wash hands thoroughly with antibacterial soap and wipe dry. 2. Put on protective clothing (e.g., sterile surgical gown, scrubs, laboratory coat). 3. Put on face-mask, protective glasses, hair cover and, if required, shoe covers. 4. Put gloves on. 5. Change gloves at least once for each new sample. 6. At the end of collection, remove and dispose of protective clothing.

EXPANDING AND FREEZING PLACENTAL FIBROBLASTS Fibroblast yield from 6- to 8-week-old placentas is usually 300,000 to 500,000 cells (PD1). To produce a population close to 10 million, cells have to divide 5 to 6 times. During this time, cells are tested for the presence of mycoplasma and several other common pathogens that can be tested in house with PCR. Only mycoplasma-negative cell lines will be frozen, usually at PD5 or 6. Testing whether the line will support growth and derivation of hESC is done in batches, usually 10 to 12 lines at once. One vial of each line is thawed, expanded up to PD8 to PD10, and mitotically inactivated with mitomycin C or γ-irradiation, and then tested for growth and maintenance of already established hESC. The cell line is considered as supportive if it maintains undifferentiated growth of the established hESC lines for 3 to 4 passages. The next step would be conducting a panel of assays recommended by the Food and Drug Administration (FDA) for producing biologicals to determine that the cells are indeed contaminant-free. These assays are done in commercial certified laboratories and since the tests are expensive, only lines considered supportive for hESC culture should be examined. Once the line is certified

BASIC PROTOCOL 2

Embryonic and Extraembryonic Stem Cells

1C.6.7 Current Protocols in Stem Cell Biology

Supplement 5

as contaminant-free, the cells can be expanded from original stock and frozen in large number of vials at higher PD, either mitotically active or inactive (γ-irradiated).

Materials Cultures of human placental fibroblasts in 25- or 75-cm2 cell culture flasks (Basic Protocol 1) Phosphate-buffered saline (PBS), Ca2+ -, Mg2+ -free (CMF-PBS; Invitrogen, cat. no. 14190-144) 0.05% Trypsin/0.53 mM EDTA (Invitrogen, cat. no. 25300-054) Fibroblast culture medium (see recipe) Fibroblast freezing medium (see recipe) Liquid nitrogen Inverted microscope 5-ml pipet 25-cm2 and 75-cm2 cell culture flasks Cryovials Additional reagents and equipment for cell counting (Support Protocol 2) and mycoplasma testing (Harlin and Gajewski, 2005) Expand placental fibroblasts 1. From cultures that are <80% confluent, aspirate medium and rinse cells with 5 ml CMF-PBS. Cell proliferation will slow down due to contact inhibition. Therefore, the cells should be subcultured before they reach 80% confluency.

2. Add 1 ml of 0.05% trypsin/EDTA per 25-cm2 flask or 3 ml per 75-cm2 flask, swirl, and wait until cells are detached and rounded-up in single-cell suspension (∼5 min at 37◦ C). To ensure that the cells are completely detached from the flask, inspect them under inverted microscope. 3. Neutralize trypsin solution using 4 to 7 ml of fibroblast culture medium, mix cells by aspirating medium into 5-ml pipet two times and take 200-µl aliquot for cell counting. 4. Resuspend 200 µl of cell suspension in 9.8 ml of diluent in a counting vial, mix by inverting 2 to 3 times and immediately count using Particle counter (see Support Protocol 2). Repeat count 3 times and record the counts. 5. Plate 250,000 to 400,000 cells per 25-cm2 flask or 750,000 to 1,200,000 cells per 75-cm2 flask with 5 or 15 ml fibroblast medium, respectively. 6. Write on each flask the preparation ID, PD, date, and the number of cells seeded into the flask. 7. After the second passage, test cells for the presence of mycoplasma (Harlin and Gajewski, 2005). If the cultures are positive, stop here and dispose of the cells in medical waste according to the institutional protocol.

Freeze placental fibroblasts 8. Follow steps 1 through 4 to remove the cells from the flask and count the cells (Support Protocol 2). Isolation of Human Placental Fibroblast Feeder Cells

When the total number of cells in culture reaches 10,000,000 to 12,000,000 (usually between PD4 and PD6), the cells should be frozen.

9. Pool all cells together and centrifuge 5 min at 700 × g, 4◦ C.

1C.6.8 Supplement 5

Current Protocols in Stem Cell Biology

Table 1C.6.1 Parameter Specifications for the Particle Counter

Setting

Value

Aperture tube size

100 µm

Kd

59.8

Upper threshold

N/A

Lower threshold

10 µm

Count mode



Metered volume

0.5 ml

Result type (Output)

Concentration/ml

Dilution factor (Output)

50 E

Printer setting

Manual short

10. Aspirate supernatant and resuspend cells to 500,000 to 1,000,000 cells/ml in fibroblast freezing medium. Distribute cell suspension 1 ml/cryovial. Label cryovials with preparation ID, PD, date, and the number of cells in the vial. 11. Place cryovials in a Styrofoam container overnight or up to 48 hr at −70◦ C. 12. Transfer vials into liquid nitrogen for long-term storage. Vials can be stored in liquid nitrogen up to 5 years without loosing growth potential.

COUNTING CELLS USING A COULTER COUNTER The most common device used for determining the number of cells per unit volume of a suspension is a counting chamber called a hemacytometer (UNIT 1C.3), since it was originally designed for performing blood cell counts. Although hemacytometers are inexpensive and easy to use, cell counts are subject to several errors: non-uniform suspensions, improper filling of chambers, failure to adopt a convention for counting cells in contact with boundary lines or each other for consistency, and statistical error of 10% to 15%. In our laboratory we routinely use a Z1-Series Coulter Counter, a fully automated instrument, which provides accuracy, speed, versatility, and excellent precision for counting cells.

SUPPORT PROTOCOL 2

Materials Diluent (Beckman Coulter) Cleaning agent (e.g., Clenz; Beckman) Cells to be counted (Basic Protocols 1 and 2) Z1 Particle Counter (Beckman Coulter) Set up the equipment 1. Prior to sample analysis using the particle counter (i.e., Beckman Coulter counter) perform the following checks: a. Check that the system is filled with diluent. b. Check that the correct aperture tube is fitted. c. Check that the diluent jar is at least 25% filled. d. Check that the waste jar is at least 50% empty. e. Check that the Instrument is powered ON. 2. Set parameters to the specifications shown in Table 1C.6.1.

Perform a background count 3. Remove vial containing the cleaning agent (i.e., Beckman Coulter Clenz) from the aperture tube using the platform release catch.

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4. To a clean vial add 10 ml of diluent and place the vial on the platform. Raise it using the platform release catch. 5. Using the data terminal press the screen access key designated FUNCTIONS. 6. Scroll until “prime aperture” is seen on the screen. 7. Press START. This procedure is used to establish a low background count at startup and between sample runs.

8. Once this step is complete, press the screen access key for SET-UP once to bring up the analysis or count screen. 9. Perform background count by pressing START. The background count should not exceed 10,000 particles per milliliter. If it is above this value, perform additional rinses with diluent by selecting FLUSH function after pressing the screen access key designated FUNCTIONS. Once FLUSH is complete, repeat background count. Repeat these steps until background count is <10,000 particles per milliliter.

10. Write down the background count.

Count the samples 11. Once the background count is complete, press the screen access key SET-UP once to access the analysis or count screen. 12. To a clean vial add 9.8 ml of diluent and 200 µl of cell suspension. Close the vial and mix content by inverting the vial 3 to 4 times. To avoid air bubbles that may lead to false counts, do not shake vial, just invert to mix the content.

13. Place the vial on the platform. Raise it using the platform release catch. 14. Press START. Count results will be displayed on the data terminal screen.

15. Mix the contents of the vial between counts. Repeat counting three times. 16. Between samples flush the system twice with 10 ml diluent added to a clean vial.

Calculate the number of cells 17. Calculate the number of cells harvested and population doubling (PD). Calculate the average of the three cell counts and record. 18. Subtract the background count. This gives the concentration of cells per milliliter of suspension.

19. Calculate the total number of cells in the tube by multiplying the average count by the total volume (in ml). These steps are summarized in the following formulas: number of harvested cells/ml = (count 1 + count 2 + count 3)/3 − background count total number of harvested cells = number of cells harvested per ml × cell suspension volume in ml Isolation of Human Placental Fibroblast Feeder Cells

If multiple flasks are harvested together, divide the total number of cells by the number of flasks to get the number of cells per flask.

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Current Protocols in Stem Cell Biology

20. Calculate the PD by dividing the average number of cells that were harvested per flask by the number of cells seeded into the flask. Then calculate the log10 of that number and divide it with the log10 2 (= 0.301). PD = [log10 (number of harvested cells /number of seeded cells)]/0.301 The electrode remains immersed in cleaning fluid.

Shut down the counter 21. To shut down particle counter, flush with 10 ml diluent. Replace the vial with a vial containing cleaning agent ensuring that the aperture and electrode are submerged, and flush again. 22. Turn particle counter OFF. The electrode remains immersed in cleaning fluid.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Collagenase solution I 100,000 U collagenase type I-A, (e.g., 0.312 g of collagenase type I-A with activity of 320 U/mg; Sigma, cat. no. C2674) 150,000 U hyaluronidase type I (e.g., 0.342 g hyaluronidase type I-S from bovine testes with activity of 439 U/mg, lyophilized; Sigma, cat. no. H3506) 120,000 Kunitz units deoxyribonuclease I (e.g., 0.060 g of deoxyribonuclease I from bovine pancreas, grade II with activity of 2,000 U/mg, lyophilized; Roche, cat. no. 104159) 0.500 g bovine serum albumin (BSA; Sigma, cat. no. A7906) 500 ml phosphate-buffered saline (PBS), Ca2+ -, Mg2+ -free (CMF-PBS; Invitrogen, cat. no. 14190-144) Add weighed enzymes and albumin to 500 ml of CMF-PBS, and stir at room temperature for 30 min or until dissolved. Filter the solution through low-proteinbinding, 0.22-µm filter unit (Corning, cat. no. 431097). Divide into 25-ml aliquots and store up to 6 months at −20◦ C. Avoid repeated freeze/thaw cycles. If the unit activity per milligram of enzyme is different than in examples, adjust weight appropriately. If <500 ml volume is desired, calculate weight appropriately. It is important that enzyme activity in the solution is as follows:

200 U/ml collagenase type I-A 300 U/ml hyaluronidase type I 240 Kunitz units/ml deoxyribonuclease I. The specific activity of a given DNase I preparation reflects the potency of the enzyme per unit mass in degrading double-stranded DNA. Historically, this activity has been expressed in Kunitz units, where 1 Kunitz unit is the amount of DNase I added to 1 mg/ml salmon sperm DNA that causes an increase of 0.001 absorbance units per min when assayed in a 0.1 M sodium acetate (pH 5.0) buffer. Embryonic and Extraembryonic Stem Cells

1C.6.11 Current Protocols in Stem Cell Biology

Supplement 5

Cytowash medium 500 ml Dulbecco’s modified Eagle medium (DMEM), high glucose (Invitrogen, cat. no. 11965-092) 5 ml Glutamine Plus (Atlanta Biological, cat. no. B90210) 12.5 ml fetal bovine serum (Hyclone, cat. no. SH30071.03) 50 mg/ml gentamicin (Invitrogen, cat. no. 15750-060) 5 ml penicillin/streptomycin (10,000 U/ml penicillin G sodium, 10,000 µg/ml streptomycin sulfate in 0.85% saline; Invitrogen, cat. no. 15140-122) Mix all components in 500 ml cellulose acetate, low-protein-binding, 0.22-µm filter unit (Corning, cat. no. 431097) and filter the mixture. Store up to 1 month at +4◦ C. Fetal bovine serum is divided in 12.5 ml-aliquots and stored up to 1 year at −20◦ C.

Fibroblast culture medium 360 ml DMEM, high glucose (Invitrogen, cat. no. 11965-092) 90 ml M199 (Invitrogen, cat. no. 11150-059) 50 ml fetal bovine serum (Hyclone, cat. no. SH30071.03) Mix all components in 500-ml cellulose acetate, low-protein-binding, 0.22-µm filter unit (Corning, cat. no. 431097) and filter the mixture. Store up to 1 month at +4◦ C. Fetal bovine serum is divided in 50-ml aliquots and stored up to 1 year at −20◦ C.

Fibroblast freezing medium 18 ml fibroblast culture medium (see recipe) 2 ml dimethyl sulfoxide (DMSO; Sigma, cat. no. D2650) Fibroblast freezing medium is 10% DMSO in fibroblast culture medium, and it is prepared by mixing DMSO with fibroblast culture medium in 1:9 ratio. The volume of the fibroblast freezing medium to be prepared depends on the total number of cells and cells per vial that will be frozen. Between 0.5 and 5 million cells can be frozen in 1 ml of freezing medium in one cryovial. Freezing medium can be used within 2 days if kept at 4◦ C.

Trypsin solution 375,000 BAEE units trypsin type I (e.g., trypsin type I with activity of 10,000 BAEE units/mg; Sigma, cat. no. T8003) 120,000 Kunitz units deoxyribonuclease I (e.g., 0.060 g of deoxyribonuclease I from bovine pancreas, grade II with activity of 2,000 U/mg lyophilizate; Roche, cat. no. 104159) 0.5 mM ethylenediamine tetraacetic (EDTA), pH 8.0 (e.g., 0.5 ml 0.5 M EDTA; Sigma, cat. no. 03690) 500 ml phosphate-buffered saline (PBS), Ca2+ -, Mg2+ -free (CMF-PBS; Invitrogen, cat. no. 14190-144) Add weighed enzymes and albumin to 500 ml of CMF-PBS, and stir at room temperature for 30 min or until dissolved. Filter the solution through low-proteinbinding, 0.22-µm filter unit (Corning, cat. no. 431097). Divide into 25-ml aliquots and store up to 6 months at −20◦ C. Avoid repetitive freezing/thawing cycles. Isolation of Human Placental Fibroblast Feeder Cells

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If the unit activity per milligram of enzyme is different than in examples, adjust weight appropriately. continued Current Protocols in Stem Cell Biology

If <500 ml volume is desired, calculate weight appropriately. It is important that enzyme activity in the solution is as follows:

750 BAEE units/ml trypsin type I 240 Kunitz units/ml deoxyribonuclease I. The specific activity of a given trypsin preparation reflects the potency of the enzyme per unit mass in hydrolysis of synthetic substrate N-α-benzoyl-L-arginine ethyl ester (BAEE), and therefore, trypsin activity is expressed in BAEE units. One BAEE unit will produce a
A253 of 0.001 per min at pH 7.6 at 25◦ C using 0.00025M (BAEE) as substrate in 3.2 ml reaction volume = 3.2 ml (1-cm light path).

COMMENTARY Background Information The establishment of the first permanent hESC line was achieved by using mouse embryonic fibroblasts (MEF) as feeder cells and fetal bovine serum (FBS)-containing culture media (Thomson et al., 1998; Reubinoff et al., 2000). The next generation of hESC culture methods was established to replace animalorigin feeder cells and FBS-containing culture media with alternatives. Several attempts were made to replace MEF with feeders of human origin. Feeders derived from human fetal muscle, fetal skin, and adult fallopian tube (Richards et al., 2002, 2003), as well as adult marrow stroma cells (Cheng et al., 2003) or uterine endometrium cells (Lee et al., 2005) supported undifferentiated growth of hESC, but not hESC derivation. Whereas at least three groups reported that immortalized fibroblastlike cells derived from hESC support growth of undifferentiated hESC (Xu et al., 2004; Stojkovic et al., 2005), only one could achieve both derivation and growth of hESC on such feeders (Wang et al., 2005). Since hESCderived feeders are a complex mixture containing uncharacterized cell types, which are isolated using laboratory-specific protocols, their ability to maintain hESCs in an undifferentiated state is likely to vary from laboratory to laboratory. The most popular MEF alternatives are human foreskin fibroblasts (HFF). Mitotically inactivated HFF between population doublings 9 and 25 have been used for both derivation of new hESC lines and continuous hESC culture (Amit et al., 2003; Hovatta et al., 2003). Human placental fibroblasts are also reported as an alternative source of human feeder cells that could support both derivation and maintenance of hESC (Genbacev et al., 2005; Simon et al., 2005). Placenta: Basic anatomy and physiology The placenta forms the fetal portion of the interface between the embryo/fetus and the

mother and it is the first organ to function during development. Human placental development depends critically on the differentiation of the placenta’s specialized epithelial cells, cytotrophoblasts (CTB, Brosens and Dixon, 1966; Boyd and Hamilton, 1967; Cross et al., 1994). Two differentiation pathways exist (Fig. 1C.6.5). In one, CTB remain in the fetal compartment and fuse to form multinucleate syncytiotrophoblasts (STB) that cover the floating chorionic villi. The primary function of STB, transport, is ideally served by their location at the villus surface. These floating villi, which are in direct contact with maternal blood in the intervillous space, perform nutrient and gas exchange for the fetus. Villous STB are covered by microvilli, multiplying severalfold the surface area. In the other differentiation pathway a subset of CTB that anchor chorionic villi also fuse, but most of them remain as single cells that detach from their basement membrane and form aggregates—cell columns—that attach to the uterine wall. From there, CTB at the distal ends of these columns attach to and then deeply invade the uterine wall (interstitial invasion) and its blood vessels (endovascular invasion) as far as the first third of the myometrium. During this process CTBs replace the endothelial and muscular lining of uterine vessels, a process that greatly enlarges the diameter of arterioles, initiates maternal blood flow to the placenta, and permits venous return to the maternal circulation. As in most organs, the placenta retains a pool of undifferentiated stem cells that are evident even at term. Whether they can compensate for placental damage by differentiating later in gestation is an interesting possibility that has been hard to prove. Cellular components of villous stroma The basic architecture of the villous stroma is a network of fibers, connective tissue cells, and fetal blood vessels. Depending on the age of placentas and the type of the villus,

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Figure 1C.6.5 Diagram of placental villi at 6 to 8 weeks of pregnancy in vivo and after dissection. (A) Fetal-maternal interface at 6 to 8 weeks. Anchoring villus (AV) functions as a bridge between fetal and maternal compartments of the human placenta. iCTB, invasive cytotrophoblasts; IVS, intervillous space; UW, uterine wall. (B) Diagram of the placental villi after dissection. Both AV and floating villi (FV) are covered with syncytiotrophoblasts (STB). Cytotrophoblasts (CTB) in anchoring villi form cell columns (CC) that attach fetal-placental unit to the uterine wall. Abbreviations: BV, blood vessels; PF, placental fibroblasts; vCTB, villous CTB; VC, villus core. Drawing: Joseph Hill, Hill-SciArt.

Isolation of Human Placental Fibroblast Feeder Cells

different types of connective tissue cells have been described (Boyd and Hamilton, 1967; Kaufmann et al., 1977; Castellucci et al., 1980; Martinoli et al., 1984; King, 1987). From our experience, cells from villous stroma at the gestational age of 6 to 8 weeks, but not from terminal placentas, can serve as feeder cells that support derivation of human embryonic stem cells (hESC; Genbacev et al., 2005; Simon et al., 2005). At this gestational age connective tissue cells in villous stroma are mostly undifferentiated mesenchymal cells and fibroblasts (Kohnen et al., 1996; Demir et al., 1997). These two types of cells can be distinguished by presence of cytoskeletal filament desmin. Whereas undifferentiated mesenchymal cells express only vimentin, fibroblasts acquire desmin as the second cytoskeletal filament (Kohnen et al., 1996, 1997). Placental fibroblasts in the villous core at this gestational age are a relatively heterogeneous population of cells with elongated cell bodies measuring 20 to 30 µm in length. From the cell bodies, often several long branching cytoplasmic processes establish contact to the extensions of neighboring cells forming a kind of reticular structure characteristic only for intermediate villi (Castellucci and Kaufmann, 1982a,b; Castellucci et al., 1984; Martinoli

et al., 1984). In later stages of pregnancy, acquisition of α-smooth muscle actin represents a first step of differentiation towards the myofibroblastic phenotype (Kohnen et al., 1996). Extracellular matrix components of villous stroma Extracellular matrix components of placental villi, such as heparan sulfate, laminin, and collagen IV, are not restricted to basement membranes but are also present throughout the stroma suggesting increasing morphogenetic and functional flexibility of the various villous cell populations (Yamada et al., 1987; Nanaev et al., 1991; Rukosuev, 1992; Muhlhauser et al., 1996). Other extracellular matrix components of placental villi include collagen I, III, IV, VI, fibronectin, tenescin, and decorin (Amenta et al., 1986; Pfister et al., 1988; Virtanen et al., 1988; Earl et al., 1990; AutioHarmainen et al., 1991; Castellucci et al., 1991; Nanaev et al., 1991; Rukosuev, 1992; Frank et al., 1994; Muhlhauser et al., 1996).

Critical Parameters Blood contamination Contamination of primary preparations with cells of nonfibroblastic origin often has a negative effect on proliferation of cells in

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culture. The most common contaminants are blood cells, and therefore, it is important to wash out as much of the blood as possible during tissue collection. If the setting is not suitable for washing the tissues at the site of collection, repeat washes in biosafety cabinet in the laboratory prior to starting the dissection of the villi. Overdigestion It is essential to check enzyme activity and calculate volumes properly prior to the preparation of enzyme stock solutions. Although tempting, we would advise against commercially available trypsin solutions (e.g., 0.25% or 0.05% trypsin/0.53 mM EDTA) widely used in tissue culture facilities if the enzyme activity is not specified in units. Inappropriate enzyme activity per volume would result in insufficient digestion (troubleshooting is easy— prolonging incubation time) or overdigestion (no troubleshooting possible; overly digested tissue has to be discarded). However, since trypsin biological activity, detachment, and disaggregation of cellular monolayer in culture dish can be monitored under the microscope and stopped any time, there is no problem in using such commercial trypsin solutions for expansion of growing cells. Contamination with pathogens Only cell lines that are negative for all pathogens should be used as feeders for hESC derivation and/or culture. Since pathogen testing by commercial certified laboratories is expensive, except for mandatory testing for mycoplasma, we initially screen all our placental fibroblast lines by PCR in house for the detection of bacteria and viruses that commonly infect the placenta.

Troubleshooting If the starting native tissue is not of a sufficient weight or if the preparation does not yield the expected number of the cells, the preparation should be discarded. Also, if there is cross-contamination between samples or contamination with bacteria, yeast, fungi, or mycoplasma, everything should be discarded. Do not try to “rescue” the sample at any cost. If the aim is to use these cells as feeders for derivation of hESC, they have to be of the highest quality or you may run a risk of wasting much more time, discovering several months later that they are not performing as expected.

feeder cells for derivation of hESC, placental fibroblast PD8 or lower passages are needed. For hESC propagation and expansion, PD12 through PD14 can be successfully used for some, but not all, cell lines. Additional PD and subcultures are possible, but biological responsiveness and function decrease as feeder cells deteriorate with each subsequent passage. Usually growth slows down and cells become senescent between PD30 and PD35, although, in some cases, they can proliferate as high as PD60.

Time Consideration The entire process, from obtaining the tissue until freezing of isolated cells, takes between 1 and 2 weeks, depending on initial number of isolated cells and their proliferation rate. Media and reagent preparation Preparation of stock media and enzymes may take 3 hr or more. This should be done in advance, never on the day of the preparation. Isolation of human placental fibroblast This is a 1-day procedure. It may take between 4 and 12 hr, depending on time spent collecting and transporting placentas, number of placentas, and experience of the person doing the preparation. Expanding and freezing placental fibroblasts Fibroblast yield from 6- to 8-week-old placentas is usually 400,000 to 500,000 cells (PD1), and the cells have to divide 5 to 6 times to produce a population close to 10 million. Since doubling time varies between 12 and 24 hr, this step may take between 6 to 12 days (this includes a lag period of 2 to 3 days after the initial plating of freshly isolated fibroblasts). Passage, counting, and freezing are not time-consuming processes. They should not take >1 hr each, regardless of the experience of the investigator.

Literature Cited Amenta, P.S., Gay, S., Vaheri, A., and MartinezHernandez, A. 1986. The extracellular matrix is an integrated unit: Ultrastructural localization of collagen types I, III, IV, V, VI, fibronectin, and laminin in human term placenta. Coll. Relat. Res. 6:125-152.

Anticipated Results

Amit, M., Margulets, V., Segev, H., Shariki, K., Laevsky, I., Coleman, R., and Itskovitz-Eldor, J. 2003. Human feeder layers for human embryonic stem cells. Biol. Reprod. 68:2150-2156.

Human placental fibroblasts isolated from 6- to 8-week-old placentas have a finite lifespan in vitro. If they are going to be used as

Autio-Harmainen, H., Sandberg, M., Pihlajaniemi, T., and Vuorio, E. 1991. Synthesis of laminin and type IV collagen by trophoblastic cells and

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fibroblastic stromal cells in the early human placenta. Lab. Invest. 64:483-491. Boyd, J.D. and Hamilton, W.J. 1967. Development and structure of the human placenta from the end of the 3rd month of gestation. J. Obstet. Gynaecol. Br. Commonw. 74:161-226. Brosens, I. and Dixon, H.G. 1966. The anatomy of the maternal side of the placenta. J. Obstet. Gynaecol. Br. Commonw. 73:357-363. Castellucci, M. and Kaufmann, P. 1982a. A threedimensional study of the normal human placental villous core: II. Stromal architecture. Placenta 3:269-285. Castellucci, M. and Kaufmann, P. 1982b. Evolution of the stroma in human chorionic villi throughout pregnancy. Bibl Anat. 22:40-45. Castellucci, M., Zaccheo, D., and Pescetto, G. 1980. A three-dimensional study of the normal human placental villous core. I. The Hofbauer cells. Cell Tissue Res. 210:235-247. Castellucci, M., Schweikhart, G., Kaufmann, P., and Zaccheo, D. 1984. The stromal architecture of the immature intermediate villus of the human placenta. Functional and clinical implications. Gynecol. Obstet. Invest. 18:95-99. Castellucci, M., Classen-Linke, I., Muhlhauser, J., Kaufmann, P., Zardi, L., and ChiquetEhrismann, R. 1991. The human placenta: A model for tenascin expression. Histochemistry 95:449-458. Cheng, L., Hammond, H., Ye, Z., Zhan, X., and Dravid, G. 2003. Human adult marrow cells support prolonged expansion of human embryonic stem cells in culture. Stem Cells 21:131-42. Cross, J.C., Werb, Z., and Fisher, S.J. 1994. Implantation and the placenta: Key pieces of the development puzzle. Science 266:1508-1518. Demir, R., Kosanke, G., Kohnen, G., Kertschanska, S., and Kaufmann, P. 1997. Classification of human placental stem villi: Review of structural and functional aspects. Microsc. Res. Tech. 38:29-41. Earl, U., Estlin, C., and Bulmer, J.N. 1990. Fibronectin and laminin in the early human placenta. Placenta 11:223-231. Frank, H.G., Malekzadeh, F., Kertschanska, S., Crescimanno, C., Castellucci, M., Lang, I., Desoye, G., and Kaufmann, P. 1994. Immunohistochemistry of two different types of placental fibrinoid. Acta Anat. (Basel) 150:55-68. Genbacev, O., Krtolica, A., Zdravkovic, T., Brunette, E., Powell, S., Nath, A., Caceres, E., McMaster, M., McDonagh, S., Li, Y., Mandalam, R., Lebkowski, J., and Fisher, S.J. 2005. Serum-free derivation of human embryonic stem cell lines on human placental fibroblast feeders. Fertil. Steril. 83:1517-1529.

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Harlin, H. and Gajewski, T.F. 2005. Diagnosis and treatment of mycoplasma-contaminated cell cultures. Curr. Protoc. Microbiol. 0:A.3B.1A.3B.6.

Hovatta, O., Mikkola, M., Gertow, K., Stromberg, A.M., Inzunza, J., Hreinsson, J., Rozell, B., Blennow, E., Andang, M., and Ahrlund-Richter, L. 2003. A culture system using human foreskin fibroblasts as feeder cells allows production of human embryonic stem cells. Hum. Reprod. 18:1404-1409. Kaufmann, P., Stark, J., and Stegner, H.E. 1977. The villous stroma of the human placenta. I. The ultrastructure of fixed connective tissue cells. Cell Tissue Res. 177:105-121. King, B.F. 1987. Ultrastructural differentiation of stromal and vascular components in early macaque placental villi. Am. J. Anat. 178:3044. Kohnen, G., Kertschanska, S., Demir, R., and Kaufmann, P. 1996. Placental villous stroma as a model system for myofibroblast differentiation. Histochem. Cell Biol. 105:415-429. Kohnen, G., Mackenzie, F., Collett, G.F., Campbell, S., Davenport, A.P., Cameron, A.D., and Cameron. I.T. 1997. Differential distribution of endothelin receptor subtypes in placentae from normal and growth-restricted pregnancies. Placenta 18:173-180. Lee, J.B., Lee, J.E., Park, J.H., Kim, S.J., Kim, M/K., Roh, S.I., and Yoon, H.S. 2005. Establishment and maintenance of human embryonic stem cell lines on human feeder cells derived from uterine endometrium under serumfree condition. Biol. Reprod. 72:42-49. Martinoli, C., Castellucci, M., Zaccheo, D., and Kaufmann, P. 1984. Scanning electron microscopy of stromal cells of human placental villi throughout pregnancy. Cell Tissue Res. 235:647-655. Muhlhauser, J., Marzioni, D., Morroni, M., Vuckovic, M., Crescimanno, C., and Castellucci, M. 1996. Codistribution of basic fibroblast growth factor and heparan sulfate proteoglycan in the growth zones of the human placenta. Cell Tissue Res. 285:101-107. Nanaev, A.K., Rukosuev, V.S., Shirinsky, V.P., Milovanov, A.P., Domogatsky, S.P., Duance, V.C., Bradbury, F.M., Yarrow, P., Gardiner, L., d’Lacey, C., and et al. 1991. Confocal and conventional immunofluorescent and immunogold electron microscopic localization of collagen types III and IV in human placenta. Placenta 12:573-595. Pfister, C., Scheuner, G., Bahn, H., and Stiller, D. 1988. Immunohistochemical demonstration of fibronectin in the human placenta. Acta Histochem. 84:83-91. Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399-404. Richards, M., Fong, C.Y., Chan, W.K., Wong, P.C., and Bongso, A. 2002. Human feeders support prolonged undifferentiated growth of human inner cell masses and embryonic stem cells. Nat. Biotechnol. 20:933-936.

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Richards, M., Tan, S., Fong, C.Y., Biswas, A., Chan, W.K., and Bongso, A. 2003. Comparative evaluation of various human feeders for prolonged undifferentiated growth of human embryonic stem cells. Stem Cells 21:546-556. Rukosuev, V.S. 1992. Immunofluorescent localization of collagen types I, III, IV, V, fibronectin, laminin, entactin, and heparan sulphate proteoglycan in human immature placenta. Experientia 48:285-287. Simon, C., Escobedo, C., Valbuena, D., Genbacev, O., Galan, A., Krtolica, A., Asensi, A., Sanchez, E., Esplugues, J., Fisher, S., and Pellicer, A. 2005. First derivation in Spain of human embryonic stem cell lines: Use of long-term cryopreserved embryos and animal-free conditions. Fertil. Steril. 83:246-249. Stojkovic, P., Lako, M., Stewart, R., Przyborski, S., Armstrong, L., Evans, J., Murdoch, A., Strachan, T., and Stojkovic, M. 2005. An autogeneic feeder cell system that efficiently supports growth of undifferentiated human embryonic stem cells. Stem Cells 23:306-314. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S.,

and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Virtanen, I., Laitinen, L., and Vartio, T. 1988. Differential expression of the extra domain-containing form of cellular fibronectin in human placentas at different stages of maturation. Histochemistry 90:25-30. Wang, Q., Fang, Z.F., Jin, F., Lu, Y., Gai, H., and Sheng, H.Z. 2005. Derivation and growing human embryonic stem cells on feeders derived from themselves. Stem Cells 23:12211227. Xu, C., Jiang, J., Sottile, V., McWhir, J., Lebkowski, J., and Carpenter, M.K. 2004. Immortalized fibroblast-like cells derived from human embryonic stem cells support undifferentiated cell growth. Stem Cells 22:972-980. Yamada, T., Isemura, M., Yamaguchi, Y., Munakata, H., Hayashi, N., and Kyogoku, M. 1987. Immunohistochemical localization of fibronectin in the human placentas at their different stages of maturation. Histochemistry 86:579584.

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Derivation of Human Skin Fibroblast Lines for Feeder Cells of Human Embryonic Stem Cells

UNIT 1C.7

Christian Unger,1 Ulrika Felldin,1 Agneta Nordenskj¨old,1 M. Sirac Dilber,1 and Outi Hovatta1 1

Karolinska Institutet, Karolinska University Hospital, Stockholm, Sweden

ABSTRACT After the first derivations of human embryonic stem cell (hESC) lines on fetal mouse feeder cell layers, the idea of using human cells instead of mouse cells as feeder cells soon arouse. Mouse cells bear a risk of microbial contamination, and nonhuman immunogenic proteins are absorbed from the feeders to hESCs. Human skin fibroblasts can be effectively used as feeder cells for hESCs. The same primary cell line, which can be safely used for up to 15 passages after stock preparations, can be expanded and used for large numbers of hESC derivations and cultures. These cells are relatively easy to handle and maintain. No animal facilities or animal work is needed. Here, we describe the derivation, culture, and cryopreservation procedures of research grade human skin fibroblast lines. We also describe how to make feeder layers for hESC using these fibrobC 2008 by John Wiley & Sons, lasts. Curr. Protoc. Stem Cell Biol. 5:1C.7.1-1C.7.10.  Inc. Keywords: human embryonic stem cells (hESCs) r human foreskin fibroblasts r derivation and expansion r freezing and thawing r feeder cell layer

INTRODUCTION Human skin fibroblast can be cultured as primary cell lines, which support the undifferentiated growth of human embryonic stem cells (hESCs) at least as effectively as mouse fetal fibroblasts. Most derived cell lines can expand for >20 passages, but due to the variability of donor tissue, a growth decline might already occur after passage 15. If a master cell bank is made by cryopreservation and vials are then expanded for use when needed, one human fibroblast cell line can be used extensively. A big advantage is that they are human cells, and all animal work can be avoided. It is possible to derive completely xeno-free feeder cell lines but, for research purposes, using fetal bovine serum (FBS) in the derivation is acceptable, and it costs much less than human serum. Here, we describe tested protocols for the derivation of research-grade human skin fibroblast feeder cells. The unit describes the simple, basic laboratory equipment and materials needed, and the culture and cryopreservation media. Protocols for the collection and transport of the skin from the operation theater are described. The derivation, propagation, and cryopreservation methods are given, as well as methods for mitotic inactivation using irradiation or mitomycin C. The unit also describes how to prepare a feeder layer for hESC.

STRATEGIC PLANNING To be able to derive primary human foreskin fibroblast cells from young boys, a special contact to a surgeon who has access to this tissue is required. For derivation, an ethics approval is required. A time plan concerning foreskin donations should be obtained Current Protocols in Stem Cell Biology 1C.7.1-1C.7.10 Published online June 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c07s5 C 2008 John Wiley & Sons, Inc. Copyright 

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and coordinated with the practical work in the laboratory. Depending on the ethical regulations in each country, one may need to obtain informed consent, which requires preparation time for the documents and time to consult parents to give the consent. Screening for major pathogens is strongly recommended if one conducts work with primary human tissue. This screening can be done preferentially from donor blood (also to be included in ethics permission) or from the final derived cell material. Until the results of pathogen screening are known, strong caution must be taken and the risks should be evaluated. The culture of primary human cells with its involved risks requires the work to be done or supervised by skilled cell culture personnel applying good laboratory practice (GLP) and good cell culture practice (GCCP; Coecke et al., 2005; Ezzelle et al., 2008). NOTE: All protocols involving the use of human materials require prior approval from the institutional ethics committee. Additionally, prior informed consent must be obtained for donated human materials. BASIC PROTOCOL

DERIVATION OF HUMAN SKIN FIBROBLAST LINES FOR FEEDER CELLS OF HUMAN EMBRYONIC STEM CELLS This protocol is used to derive human foreskin fibroblast cells for research-based human embryonic stem cell culture. These cells can usually be grown for 15 or more passages and provide a human feeder alternative to mouse embryonic fibroblasts. Although applicable for longer periods and easier to grow (without gelatin), their human character requires a higher degree of strategic planning for their derivation (see above). Pathogen screening for mycoplasma, HIV, Hepatitis, or other relevant infections should be complete before a cell line is used as feeders. NOTE: All solutions and equipment coming in contact with living cells must be sterile and aseptic technique should be used accordingly. NOTE: All media and solutions should be at room temperature unless otherwise specified. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator.

Materials Human foreskin tissue from young boys (not older than 2 years), a few mm3 Disinfectant soap Phosphate-buffered saline (D-PBS) plus antibiotics (see recipe) Blood sample from the donor for basic pathogen screening Heat-inactivated fetal bovine serum (FBS; Invitrogen, cat. no. 10500-064 or Hyclone, cat. no. SH30070) Derivation/culture medium (see recipe) D-PBS without calcium and magnesium (Invitrogen, cat. no. 14190) TrypLE Express (Invitrogen) Trypan blue (Invitrogen)

Human Skin Fibroblast Feeder Cells

Sterile transportation tube Laminar flow hood (Class II biosafety cabinet) BD Primaria 6-well-plates (BD Biosciences) or easy grip cell culture dish with Primaria surface treatment (BD) 10-cm petri dish Dissecting forceps and fine scissors, sterilized by autoclaving 12-ml tube (i.e., BD Falcon) Inverted microscope

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Hemacytometer (e.g., from Marienfeld GmbH, Germany) Tissue culture–treated plasticware: 6-well plates, 25-cm2 and 75-cm2 flasks Additional reagents and equipment for counting viable cells using a hemacytometer and trypan blue staining (UNIT 1C.3) CAUTION: Take special care when working with human primary tissue!

Collect tissue The following steps are performed by a surgeon in an operating theater. 1. Wash donor skin thoroughly preoperatively with disinfectant soap. 2. Have the surgeon cut skin from the penile shaft or the prepuce. 3. Place the skin (>10-mm3 ) into a sterile transportation tube with sterile salt solution or D-PBS plus antibiotics. 4. Transport tissue pieces from operation theater (plus blood sample for viral tests, if possible) to cell culture laboratory in sterile container as quickly as possible. Transport and waiting periods >12 hr are not recommended.

Isolate fibroblasts For aseptic work conditions, all further steps involving cell manipulation must be done in a laminar flow hood. 5. Cover Primaria 6-well plate with 0.5 ml/well freshly thawed FBS, aspirate, and let the remaining FBS dry for 10 min with open lid before seeding tissue on the plate. 6. Take tissue to a sterile working environment and place onto a 10-cm petri dish. 7. Cut the tissue, using sterile forceps and scissors, into small pieces not larger than ∼1 mm3 . 8. Resuspend the tissue pieces in 6 ml derivation/culture medium, onto 2 wells of a serum-coated Primaria 6-well plate (3 ml tissue solution per well; 1 well of a 6-well plate is ∼10 cm2 in size) and leave undisturbed until first fibroblasts attach. 9. Check the plates after 1 to 2 days to see if tissue is contaminated by bacteria and re-check the results of the pathogen screening from the donor blood (if available). To detect bacterial infections no antibiotics are added to the derivation/culture medium. Do NOT continue with infected material!

10. Add 3 ml fresh derivation/culture medium per well if the medium is changing pH (color indicator), but latest after 3 days. 11. Remove ∼2/3 of the medium on day 6 after seeding, put it into a 12-ml tube, and let non-attached cell clumps settle for 2 min by gravitation. This step removes dead cells and debris from the potential outgrowing cells and adds new nutrition.

12. Replace the used medium with 2 ml fresh derivation/culture medium and put resuspended tissue clumps back in the same well with the already attached cells. 13. Continue cell culture by changing medium every 4 to 5 days, or if the pH changes, until the fibroblast cells (Fig. 1C.7.1) grow to confluence. Settle non-attached tissue-clumps as in step 11, as long as few fibroblast cells have attached and the culture well is not yet confluent

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Figure 1C.7.1 Phase contrast picture of confluent primary human foreskin fibroblast cells (A) at low (4×) and (B) higher (10×) magnification.

Propagate human primary foreskin fibroblast cells It is necessary to subculture the cells to achieve an appropriate amount of cells, depending on the purpose. For vital freezing, use no less than 1 × 106 cells/vial, preferably at a low passage. 14. Remove old medium from the cells. 15. Wash three times, each time with 4 ml D-PBS to remove any residual FBS that may inhibit the action of TrypLE Express. 16. Remove all D-PBS with a pipet. 17. Add an appropriate amount of TrypLE Express to cover the cell layer. Use 0.5 ml/well in a 6-well plate, 0.7 ml/25-cm2 flask, or 2 ml/75-cm2 flask.

18. Incubate the cells until they detach, ∼5 to 10 min. To confirm that the cells have detached, tap the flask carefully and check under a microscope that the cells have detached.

19. Add at least 2 ml of derivation/culture medium per 0.5 ml TrypLE, then rinse the cell layer three times to dissociate cells and to dislodge any remaining adherent cells.

Determine the cell number 20. Transfer the cell suspension to a 12-ml tube and measure the exact volume so you will be able to calculate the total cell number. 21. Count the number of viable cells (i.e., using standard trypan blue staining and hemacytometer; UNIT 1C.3). Cell density at confluence is 1 to 2 × 106 /75-cm2 flask depending on the size of the cells.

22. Seed 0.5 × 106 cells in 13 ml derivation/culture medium in a 75-cm2 cell culture flask, or use a flask that is suitable for the number of cells obtained. Always use a new flask otherwise the cells will not attach properly. Cultures should be labeled with passage number, date, and signature. It is important that the cells have cell-cell contact, otherwise they will not expand properly. After the first passaging with trypsin, cells are labeled passage 1.

23. Passage culture when it becomes confluent by repeating steps 14 to 22. SUPPORT PROTOCOL 1 Human Skin Fibroblast Feeder Cells

VITAL FREEZING, THAWING, AND THE RECOVERY OF FEEDER CELLS This protocol describes vital freezing, thawing, and the recovery of feeder cells. For long-term storage and future use it is necessary to obtain a stock of cells frozen at an early passage (≤passage 3).

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Materials Early passage (≤3) human foreskin fibroblasts (Basic Protocol) Heat-inactivated fetal bovine serum, ice cold Freezing medium (see recipe), ice cold 70% ethanol Derivation/culture medium (see recipe) 12-ml tubes Centrifuge with swing-out rotor, able to spin 12-ml tubes 2-ml cryovials Freezing container (Styrofoam box or Mr. Frosty; Nalgene) −80◦ C freezer −150◦ C freezer or nitrogen tank 37◦ C water bath 75-cm2 culture flask Additional reagents and equipment for washing and detaching the cells (Basic Protocol) Perform vital freezing 1. Wash and detach the cells as described in Basic Protocol, steps 14 to 22. Collect the cells in a 12-ml tube. 2. Centrifuge the cells 7 min at 200 × g, room temperature, and discard the supernatant. 3. Resuspend the cell pellet in cold FBS to a density of 2 × 106 cells/ml. Final concentration will then be 1 × 106 cells/ml when freezing medium is added.

4. Incubate on ice for 20 min. 5. Prepare cryovials by labeling with cell type, passage number, date, and signature. 6. Add equal amount of cold freezing medium to the cell suspension, mix, and transfer 1 ml to each of the labeled 2-ml cryovials. Final DMSO concentration will be 6% and the final cell concentration 1 × 106 cells/ml.

7. Immediately put the tubes into a freezing container and move to a −80◦ C freezer to allow slow freezing (∼1◦ C/min cooling rate). 8. After 24 hr, move the tubes to a −150◦ C freezer or nitrogen tank for long-term storage. Alternatively, freezing tanks with freezing carousel or programmable freezer can be used in steps 7 and 8.

Thaw and recover cells 9. Obtain a cryovial from the freezer. 10. Thaw cells rapidly in a 37◦ C water bath by agitating tube in the water until ∼3/4 of the contents is thawed. 11. Remove the tube from the water bath but continue to agitate until the cells are completely thawed. 12. Disinfect the tube by rinsing with 70% ethanol. 13. Immediately transfer thawed cells to a 75-cm2 culture flask containing 20 ml derivation/culture medium. 14. Culture the cells until the next day.

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15. Remove old medium and add 12 ml fresh derivation/culture medium. 16. Continue with subculturing and expansion as described above (steps 14 to 23 of the Basic Protocol). SUPPORT PROTOCOL 2

MITOTIC INACTIVATION BY γ -IRRADIATION TO PREPARE HUMAN FIBROBLAST FEEDER CELLS FOR hESC CULTURE For hESC cultures ∼30,000 fibroblasts/cm2 are required to cover the tissue culture surface, and they must be mitotically inactivated. This prevents the expansion of the foreskin fibroblasts in the mixed culture with hESCs, while ensuring their supportive/feeding role. Mitotic inactivation is commonly achieved by exposure to γ-irradiation or mitomycin C treatment (see Support Protocol 3). γ-Irradiation is the preferred method if an irradiation source is available, because it avoids the use of toxic substances.

Materials Confluent cultures of human foreskin fibroblasts (Basic Protocol) Derivation/culture medium (see recipe) γ source (if available; e.g. Cs-137 irradiator, Gammacell 2000) 6-well plates

Additional reagents and equipment for washing and detaching the cells (Basic Protocol) 1. Wash and detach cells as described in Basic Protocol, steps 14 to 22. 2. Dilute the cells in derivation/culture medium to a concentration of 1.5 × 105 cells/ml. 3. Expose the cells to a total dose of 40 Gy from a controlled γ-radiation source. 4. Seed the cells in 6-well plates: 2 ml cell suspension/well, 3 × 105 cells/well final concentration. 5. Keep the cells in an incubator until further use. Prepared irradiated cells can be used the next day or up to 4 weeks after inactivation. If not used immediately, the medium should be changed weekly. SUPPORT PROTOCOL 3

MITOTIC INACTIVATION BY MITOMYCIN C TREATMENT TO PREPARE HUMAN FIBROBLAST FEEDER CELLS FOR hESC CULTURE Cells may be inactivated by mitomycin C treatment if a γ-radiation source is not available. Although this method is more time and labor intensive, the inactivated feeders are equally suitable for ES cell culture. Care must be taken due to the toxic character of mitomycin C, and the fibroblasts must be extensively washed to prevent effects on co-cultured hESCs. Refer to Conner (2001) and Nieto et al. (2007) for more detailed reading.

Materials Cultures of human skin fibroblasts (see Basic Protocol) Derivation/culture medium (see recipe) Mitomycin C solution (see recipe) D-PBS without calcium and magnesium (Invitrogen, cat. no.14190) TrypLE Express (Invitrogen) Trypan blue (Invitrogen) Human Skin Fibroblast Feeder Cells

12-ml tube Hemacytometer 6-well plates

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Additional reagents and equipment for viable cell counting using a hemacytometer and trypan blue (UNIT 1C.3) CAUTION: Mitomycin C is toxic. 1. Replace the standard derivation/culture medium on the required amount of fibroblasts with derivation/culture medium containing 10 μg/ml mitomycin C solution. 2. Incubate cells for 2.5 hr in a 37◦ C, 5% CO2 incubator 3. Discard mitomycin C–containing medium and wash cells three times, each time with 10 ml D-PBS per 75-cm2 flask. Mitomycin C–containing medium should be disposed of as hazardous waste, according to institutional health and safety guidelines.

4. Discard the washing solution and add 2 ml TrypLE Express to the 75-cm2 flask and incubate for 3 min at 37◦ C to dislodge the cells. 5. Remove the TrypLE Express with a pipet. To confirm that the cells have detached, tap the flask carefully and check under a microscope.

6. Resuspend cells in 5 ml derivation/culture medium and transfer to a 12-ml tube. 7. Count cells as in step 21 of the Basic Protocol. 8. Seed the cells in 6-well plates: 2 ml cell suspension/well, 3 × 105 cells/well final concentration.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Cell culture and derivation medium (Derivation/culture medium) Iscove’s Modified Dulbecco’s Medium with GlutaMAX (IMDM; Invitrogen, cat. no. 31980-048) 20% (v/v) freshly thawed and heat-inactivated fetal bovine serum (FBS) 1× MEM nonessential amino acids (100× stock; Invitrogen) D-PBS plus antibiotics D-PBS without calcium and magnesium (Invitrogen, cat. no. 14190) 1% penicillin-streptomycin (100× stock; Invitrogen) Store up to 1 week at −20◦ C Freezing medium 12% (v/v) dimethyl sulfoxide (DMSO) in D-PBS without calcium and magnesium (Invitrogen, cat. no. 14190) Store up to 1 month at 4◦ C CAUTION: DMSO is toxic. Use with care.

Mitomycin C solution Dissolve 2 mg mitomycin C powder (Sigma-Aldrich) in 4 ml sterile water and mix to dissolve (stock solution: 0.5 mg/ml). Sterile filter the stock solution and store for up to 2 weeks at 4◦ C. Use at a final concentration of 10 μg/ml. CAUTION: Mitomycin C is toxic.

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COMMENTARY Background Information Shortly after the advent of hESC lines, a need for more human adaptation was pushing the development of human feeder cells for hESC growth, turning away from the commonly used murine embryonic fibroblast feeder cells (Amit et al., 2003; Hovatta et al., 2003; Richards et al., 2003). We began to use human feeder cells in our laboratory in 2002 in order to reduce the number of factors potentially contaminating valuable and unique hESC lines (Hovatta et al., 2003). The use of human materials is particularly important in the derivation and culture of clinical-grade hESC lines, but at the same time it is important to carry out the research preceding clinical applications using similarly cultured lines. Various human cell types have been used, and they have been shown to support hESC growth and even their derivation. Fibroblasts supporting hESC lines have been derived from fetal skin and muscle of aborted embryos (Richards et al., 2002; Sidhu et al., 2006), and from adult tissues, such as fallopian tube and foreskin (Amit et al., 2003; Hovatta et al., 2003). Cheng et al. (2003) used human marrow-derived stromal cells to expand hESCs. Uterine endometrial cells have been used as feeders (Lee et al., 2005). Also, hESC-derived fibroblast feeders have been used (Stojkovic et al., 2005). When using such feeder cells, a hESC line has to be derived first, which requires feeder or matrix support. We have applied human foreskin fibroblasts in hESC cultures (Hovatta et al., 2003) from the first derivations, and have since then derived many hESC lines, showing their applicability (Inzunza et al., 2005; Strom et al., 2007).

Critical Parameters and Troubleshooting

Human Skin Fibroblast Feeder Cells

As noted in the beginning of this unit, it should be emphasized that all aspects of cell culture and all involved materials and equipment must be aseptic. Good laboratory practice (GLP; Ezzelle et al., 2008) and good cell culture practice (GCCP; Coecke et al., 2005) needs to be applied in order to obtain optimal results, and to minimize the risks associated with handling of human materials. A careful risk evaluation should be done with a senior researcher and the clinician involved. It is important to evaluate which infections the skin donor might carry, and how to ensure that these can be detected. The practitioner should be skilled in cell culture tech-

niques since potentially infected human skin is handled. Tissue for this protocol should only be derived from healthy donor skin. However, an extra screening for pathogens may be required to confirm that there are no major infections. A donor blood screening for common infections like mycoplasma, HIV, and hepatitis should be discussed when planning the derivation of cells and conducted when possible. If no blood can be obtained a screening of the derived cells by PCR (for example) might be considered. All these procedures can reduce the risk for you or anyone working with the involved cells (fibroblasts or hESCs). If any contamination, either from the donor or improper handling procedures, is suspected, the cells and solutions should be discarded. As all human materials, such as cell lines, should be effectively destroyed/lysed before discarding them, no additional special procedure is required. However, special procedures may be necessary if certain infections are possible and should be carefully discussed in the risk assessment. Although the procedures described here have been found to be reliable, certain factors are of importance and can influence success. The tissue needs to be cut up extensively to obtain small pieces (∼1 mm3 ). Larger tissue clumps make it difficult for fibroblasts to attach and grow out. Single cells do not expand and cell-cell contact is required to get cells growing; therefore, clumps are required. Cells will usually grow out in a layer starting from the aggregates. Cells can be grown to confluence. Then, when propagating the cells, a seeding density that provides cell-cell contact should be used. A split ratio of 1:3 has been found to create a good density after the cells have reached confluence. When the human tissue is seeded, cell types such as blood cells might be growing and using the medium. This might be causing the medium to be used up faster at the beginning of derivation. This is why one should add medium as indicated in the protocol. The subsequent complete medium changes will remove non-attached cells while the fibroblast cell layer is growing out. During the first week after seeding the cells, the main task is to check the cells and to make sure that the medium is renewed, and the cells are free of contamination. A major factor for the fibroblasts to attach and grow out is the use of serum within the medium and on the precoated surface. Serum that has long been stored at 4◦ C can lose its properties,

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and therefore the use of freshly thawed serum is strongly recommended to improve attachment. After the primary cells have started to expand and a cell layer has been formed, future passaging does not require any precoating of cell culture dishes. This is an advantage compared to the required gelatin coating for mouse embryonic fibroblast (MEF) feeders. It is useful to always passage the cells into new flasks. This ensures a more equal distribution of cells after seeding. It also puts selective pressure to expand only easily attaching cells, such as fibroblasts, which produce their own matrix proteins. This results in a decline of the possibility of other cell types initially attaching under the rich medium conditions. The numerous fibroblast expansions done in our laboratory have resulted in a cell density that is commonly reached when human foreskin fibroblasts are confluent. At this stage, there are 12,000 to 15,000 cells per cm2 . If this cell density is very different for your derived cells, maybe another cell type has been expanded. In such cases, the morphology should be carefully checked (see Fig. 1C.7.1) and the support for hESC growth should be evaluated. Human fibroblast feeder (HFF) cells can be passaged and undergo >50 population doublings, while they have been reported supporting hESC cultures up to passage 25 (Hovatta et al., 2003). At higher passages, fewer cells divide and instead the cells expand in size. By following the cell density, this transition can be noticed. Such big cells are inferior for hESC support and need to be discarded. The long life span of your human feeder stock is likely dependent on the donor age, which makes it preferential to use foreskin from young boys (preferably newborn). We have so far successfully derived HF cells from boys up to 2 years of age. A frozen stock should be prepared as early as a good number of cells has been obtained, but leaving enough life span to the cells to be used for elongated hESC culture. We commonly prepare an early frozen stock at passage 3. Finally, to use HFF cells for hESC culture, the cells need to be mitotically inactivated as indicated in the support protocols. In the case of γ-irradiation, the irradiation parameter must be checked carefully to be sure that a correct dose is applied. At doses too low, the cells might still expand and later contaminate the hESC culture. At doses too high, the cells will go directly into apoptosis and will not provide any feeder capabilities. Irradiation indicators, such as stickers on the tube, can be used to verify that the irradiation has been applied.

It needs to be considered that different tube sizes and volume of media with fibroblasts will require different irradiation times. This should be discussed with staff responsible for the irradiation facilities. If it is noticed that the cells are still partially expanding, the irradiation dose can be increased up to 60 Gy without problem; however, the effect on the individual batch of cells should be evaluated and their support of hESCs verified. Similarly, if mitomycin C is used, the solution should not be expired. Fresh solution should be prepared if in doubt. If required for proper inactivation, incubation times can be increased to 4 hr (Nieto et al., 2007).

Anticipated Results The derivation of human fibroblast feeder cells is relatively straightforward using this method. However, the need for human donors makes it necessary to plan the process carefully. Since human materials are subject to a certain donor variability, a successful outgrowth and stock preparation of human fibroblast cells is expected from 2 out of 3 tissue samples. Therefore, it is strongly recommended to obtain several foreskin tissue pieces (at least 2) to make sure to have at least one final feeder cell line. Parallel growth of cells is not a big additional effort and handling three or more derivations at the same time is possible. If cells expand successfully, an early passage 3 stock yields ∼3 to 4 vials of frozen cells with 1 × 106 cells/vial and at passage 4, 9 to 12 vials could be frozen. Fibroblast density at confluence is estimated in a range of 12,000 to 15,000 cells/cm2 . Since the cells are expected to support hESC culture for at least another 12 passages, it is a standard practice to prepare an additional working stock at passage 6/7. Although each feeder cell line is expected to support hESC growth, the available hESC lines should be successfully cultured for at least 4 passages before they are commonly used.

Time Consideration The time for planning and obtaining human foreskin tissue can vary considerably depending on the availability of donors (see Strategic Planning). Time required from derivation to original stock freezing of human foreskin fibroblast cells can be estimated from 1 to 1.5 month if stock is frozen at passage 2 to 3. This depends on tissue and preparation variability. Cells can be mitotically inactivated and seeded for human embryonic stem cell culture approximately at the same time, as the frozen stock is prepared.

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Acknowledgment This work has been supported by the European Union (ESTOOLS).

Literature Cited Amit, M., Margulets, V., Segev, H., Shariki, K., Laevsky, I., Coleman, R., and Itskovitz-Eldor, J. 2003. Human feeder layers for human embryonic stem cells. Biol. Reprod. 68:21502156. Cheng, L., Hammond, H., Ye, Z., Zhan, X., and Dravid, G. 2003. Human adult marrow cells support prolonged expansion of human embryonic stem cells in culture. Stem Cells 21:131-142. Coecke, S., Balls, M., Bowe, G., Davis, J., Gstraunthaler, G., Hartung, T., Hay, R., Merten, O.W., Price, A., Schechtman, L., Stacey, G., and Stokes, W. 2005. Guidance on good cell culture practice. A report of the second ECVAM task force on good cell culture practice. Altern. Lab. Anim. 33:261-287. Conner, D.A. 2001. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Molec. Biol. 51:23.2.1-23.2.7. Ezzelle, J., Rodriguez-Chavez, I.R., Darden, J.M., Stirewalt, M., Kunwar, N., Hitchcock, R., Walter, T., and D’souza, M.P. 2008. Guidelines on good clinical laboratory practice: Bridging operations between research and clinical research laboratories. J. Pharm. Biomed. Anal. 46:18-29. Hovatta, O., Mikkola, M., Gertow, K., Stromberg, A.M., Inzunza, J., Hreinsson, J., Rozell, B., Blennow, E., Andang, M., and Ahrlund-Richter, L. 2003. A culture system using human foreskin fibroblasts as feeder cells allows production of human embryonic stem cells. Hum. Reprod. 18:1404-1409. Inzunza, J., Gertow, K., Stromberg, M.A., Matilainen, E., Blennow, E., Skottman, H., Wolbank, S., Ahrlund-Richter, L., and Hovatta, O. 2005. Derivation of human embryonic stem

cell lines in serum replacement medium using postnatal human fibroblasts as feeder cells. Stem Cells 23:544-549. Lee, J.B., Lee, J.E., Park, J.H., Kim, S.J., Kim, M.K., Roh, S.I., and Yoon, H.S. 2005. Establishment and maintenance of human embryonic stem cell lines on human feeder cells derived from uterine endometrium under serumfree condition. Biol. Reprod. 72:42-49. Nieto, A., Cabrera, C.M., Catalina, P., Cobo, F., Barnie, A., Cortes, J.L., Barroso Del Jesus, A., Montes, R., and Concha, A. 2007. Effect of mitomycin-C on human foreskin fibroblasts used as feeders in human embryonic stem cells: Immunocytochemistry MIB1 score and DNA ploidy and apoptosis evaluated by flow cytometry. Cell Biol. Int. 31:269-278. Richards, M., Fong, C.Y., Chan, W.K., Wong, P.C., and Bongso, A. 2002. Human feeders support prolonged undifferentiated growth of human inner cell masses and embryonic stem cells. Nat. Biotechnol. 20:933-936. Richards, M., Tan, S., Fong, C.Y., Biswas, A., Chan, W.K., and Bongso, A. 2003. Comparative evaluation of various human feeders for prolonged undifferentiated growth of human embryonic stem cells. Stem Cells 21:546-556. Sidhu, K.S., Lie, K.H., and Tuch, B.E. 2006. Transgenic human fetal fibroblasts as feeder layer for human embryonic stem cell lineage selection. Stem Cells Dev. 15:741-747. Stojkovic, P., Lako, M., Stewart, R., Przyborski, S., Armstrong, L., Evans, J., Murdoch, A., Strachan, T., and Stojkovic, M. 2005. An autogeneic feeder cell system that efficiently supports growth of undifferentiated human embryonic stem cells. Stem Cells 23:306-314. Strom, S., Inzunza, J., Grinnemo, K.H., Holmberg, K., Matilainen, E., Stromberg, A.M., Blennow, E., and Hovatta, O. 2007. Mechanical isolation of the inner cell mass is effective in derivation of new human embryonic stem cell lines. Hum. Reprod. 22:3051-3058.

Human Skin Fibroblast Feeder Cells

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Cryopreservation of Dissociated Human Embryonic Stem Cells in the Presence of ROCK Inhibitor

UNIT 1C.8

˜ 1, 2 Anne Marie Str¨omberg,2 Outi Hovatta,2 and Raquel Mart´ın-Ib´anez, 1 Josep M. Canals 1

Departament de Biologia Cellular, Immunologia i Neurosci`encies, Facultad de Medicina, Institut d’Investigacions Biom`ediques August Pi i Sunyer (IDIBAPS), Universitat de Barcelona and Centro de Investigaci´on Biom´edica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Barcelona, Spain 2 Department of Clinical Science, Intervention and Technology, Karolinska Institutet, Karolinska University Hospital Huddinge, Stockholm, Sweden

ABSTRACT Two different methods have been adopted for the cryopreservation of human embryonic stem cells (hESCs): vitrification and conventional slow freezing/rapid thawing. However, these methods present poor viability and high differentiation rates. Therefore, the development of an efficient cryopreservation protocol for hESCs is one of the major challenges for the application of these cells in clinical therapy and regenerative medicine. A novel method for the cryopreservation of dissociated hESCs in the presence of a selective Rho-associated kinase (ROCK) inhibitor that increases cell survival and the efficiency of colony formation of cryopreserved hESCs has been developed. Moreover, this protocol improves the existing methods presenting short recovery times and hardly any differentiation rates. Thus, an easy handling protocol that allows the cryopreservation of large C 2009 amounts of hESCs is described. Curr. Protoc. Stem Cell Biol. 10:1C.8.1-1C.8.15.  by John Wiley & Sons, Inc. Keywords: freezing r thawing r Y-27632 r survival r differentiation r hESCs

INTRODUCTION Human embryonic stem cells (hESCs) are generally passaged and cryopreserved as small aggregates because they present a high susceptibility to apoptosis upon cellular detachment and dissociation. Thus, splitting and freezing processes become tedious and clearly unsuited for handling large amounts of cells. Consequently, it is necessary to improve the existing techniques for cryopreservation, manipulation, and handling of hESCs. In fact, it has been reported that treatment with a selective Rho-associated kinase (ROCK) inhibitor in routine enzymatic passaging procedures increases the survival of dissociated hESCs and their cloning efficiency. In accordance, the protocol described here goes one step forward in the cryopreservation of dissociated hESCs using the conventional slow freezing/rapid thawing method. A selective Rho-associated kinase (ROCK) inhibitor, Y-27632, is used to reduce cell death, thereby increasing cloning efficiency. Y-27632 is added in three critical steps of the cryopreservation protocol: the dissociation step of hESC colonies into single cells, the freezing step, and the thawing/seeding final step. This inhibitor allows the cryopreservation of large amounts of single hESCs with high survival rates and efficiency of colony formation. In addition, this protocol presents the advantage that no specific requirements are necessary to put it into practice. The effects of ROCK inhibitor on routine enzymatic passaging of hESCs have been investigated and are described in Martin-Iba˜nez et al. (2008). In this study, it has been observed that treatment of hESC colonies with the ROCK inhibitor Y-27632, before and after splitting,

Current Protocols in Stem Cell Biology 1C.8.1-1C.8.15 Published online July 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c08s10 C 2009 John Wiley & Sons, Inc. Copyright 

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mitotic inactivation

human foreskin fibroblasts feeder layer preparation (see Support Protocol 2) human foreskin fibroblasts 2 days pieces of hESC colonies 5–7 days

hESCs maintenance culture mechanical splitting (see Support Protocol 1)

mechanical splitting

5–7 days

ROCK inhibitor (1 hr) dissociation of hESC colonies

expansion of hESCs by single cell dissociation seeding of dissociated hESCs onto a feeder layer

ROCK inhibitor (1st day) 6–8 days

ROCK inhibitor (1 hr)

suspension of dissociated hESCs in freezing medium with ROCK inhibitor

freezing dissociated hESCs

freezing dissociated hESCs in a freezing container (~1°C/min) introduced in a 80°C freezer

hESC

cryopreservation of dissociated hESCs in the presence of ROCK inhibitor (see Basic Protocol)

dissociation of hESC colonies

24 hr

N2 tank thawing dissociated hESCs in a 37°C water bath

seeding of dissociated hESCs onto a feeder layer

thawing dissociated hESCs

ROCK inhibitor (1 day) 6–8 days

Figure 1C.8.1 Graphic representation of the protocols described in this unit including: human foreskin Þbroblasts feeder layer preparation, hESC maintenance culture, and cryopreservation of dissociated hESCs in the presence of ROCK inhibitor.

Cryopreservation of Dissociated hESCs

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significantly increased the efficiency of colony formation of dissociated hESCs compared to the untreated control condition—treatment with Y-27632: 5% to 7% efficiency of colony formation; untreated control: 0.02% to 0.09% efficiency of colony formation. This unit describes a protocol used for the propagation and cryopreservation of dissociated hESCs cultured on a monolayer of feeder cells. Culture and cryopreservation media are described. Basic laboratory equipment and materials needed are detailed. Finally, the unit also describes how to culture hESCs in maintenance conditions and how to prepare a feeder layer of human foreskin fibroblasts (see overall scheme of the protocol in Fig. 1C.8.1).

STRATEGIC PLANNING hESC culture on a feeder cell monolayer of human skin fibroblasts requires planning for the number of inactivated feeder cell plates that will be needed to expand or thaw hESCs. Therefore, 2 to 3 days before starting experiments with hESCs, it will be necessary to prepare the correct number of inactivated feeder cell plates. hESC culture, expansion, and cryopreservation require prior approval from the institutional ethics committee and from the corresponding national organization.

CRYOPRESERVATION OF DISSOCIATED HUMAN EMBRYONIC STEM CELLS IN THE PRESENCE OF ROCK INHIBITOR

BASIC PROTOCOL

This protocol describes how to expand and cryopreserve dissociated hESCs in the presence of a selective ROCK inhibitor, Y-27632. The addition of this inhibitor before and after dissociation allows expansion of single hESCs seeded on feeder cells. These cells give rise to a large number of colonies that are ready to be cryopreserved between 5 and 7 days. Moreover, the presence of Y-27632 at different steps of the cryopreservation protocol allows dissociated hESCs to be frozen and thawed with high survival rates, a shorter time of recovery, and almost no differentiation. NOTE: All solutions and equipment in contact with living cells must be sterile and aseptic techniques should be used accordingly. NOTE: All the materials used for cell culture should be sterilized with 70% ethanol before being introduced into the laminar flow hood. NOTE: All media and solutions should be at 37◦ C unless otherwise specified. NOTE: All culture incubations should be performed in a 37◦ C/5% CO2 humidified incubator. CAUTION: hESCs are very sensitive to changes in temperature and CO2 levels. They should be maintained out of the incubator for as short a time as possible. Do not open and close the incubator often, to avoid changes in temperature and CO2 levels.

Materials hESCs 5 mM ROCK inhibitor Y-27632 (Calbiochem, cat. no. 688001) Dulbecco’s phosphate buffered saline without calcium and magnesium (CMF-DPBS; Invitrogen, cat. no. 14190) TrypLE Express (Invitrogen, cat. no. 12604021) hESC culture medium (see recipe) Basic fibroblast growth factor (bFGF; R&D Systems, cat. no. 234-FSE/CF) Trypan blue

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Mitotically inactivated human foreskin fibroblast feeders (see Support Protocol 2) DMSO 70% ethanol Stereomicroscope Sterile 15-ml conical tubes (e.g., BD Falcon) 35-mm cell culture dishes (BD Falcon, cat. no. 353001) 200-μl micropipets Centrifuge Neubauer counting chamber or hemacytometer 1.8-ml cryotubes (Nunc, cat. no. 377267) Cryo 1◦ C freezing container (e.g., Mr. Frosty; Nalgene, cat. no. 5100) −80◦ C freezer −150◦ C freezer or liquid nitrogen tank 37◦ C water bath NOTE: ROCK inhibitor stock solution should be stored in small aliquots at −20◦ C to avoid repeated freeze/thaw cycles and also should be protected from light. NOTE: To work in sterile conditions, all steps involving cell manipulation must be done in a laminar flow hood.

Preincubate hESC cultures in ROCK inhibitor For long-term storage and further use, it is necessary to obtain a large stock of frozen hESCs, especially at early passages. Therefore, before cryopreserving hESCs, it is necessary to expand them by dissociation. 1. Check, with a stereomicroscope, the appearance of the hESC colonies that are going to be expanded. These colonies have been growing in maintenance conditions (see Support Protocol 1) for up to 6 to 8 days on a feeder layer, and should be of the right size (between 1 and 2 mm in diameter) to be split. hESC colonies to be expanded should present undifferentiated morphological features, such as well-defined borders and small cells with high nuclear:cytoplasm ratio, which can be observed under an inverted microscope (Fig. 1C.8.2).

2. If the cells present good morphological features, add 10 μM ROCK inhibitor Y-27632 stock solution to the culture medium and incubate for 1 hr. 3. After the 1-hr incubation, remove culture medium from the dish using a 1-ml pipet.

Dissociate hESCs cultures into single cells 4. Wash two times with CMF-DPBS (1 ml/35-mm dish). 5. Treat cells with TrypLE Express (0.5 ml TrypLE Express/35-mm dish) and incubate 5 min. TrypLE Express treatment is intended to dissociate only hESCs colonies but not feeders.

6. Before feeders start to detach from the plate, dilute TrypLE Express with 1 ml CMF-DPBS.

Cryopreservation of Dissociated hESCs

7. Dissociate ONLY hESC colonies by flushing TrypLE Express solution over each colony several times with a 200-μl micropipet. Avoid using more than 200 μl of the solution to flush the hESC colonies, otherwise feeders start to detach from the plate.

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A

B

Figure 1C.8.2 Bright-Þeld (A) and phase-contrast (B) photographs showing the morphological features of undifferentiated hESC colonies. Scale bar = 500 and 30 μm for low and high magniÞcation, respectively.

Figure 1C.8.3 500 μm.

Bright-Þeld photograph showing differentiated hESC colonies. Scale bar =

If some colonies of the plate present differentiated areas, do not dissociate them. Once hESC colonies with good morphological features have been dissociated and transferred to a 15-ml conical sterile tube, differentiated hESC colonies and feeders will remain on the dish. For an example of differentiated colonies, see Figure 1C.8.3. If TrypLE Express works correctly, dissociation of hESC colonies into single cells should be feasible while at the same time keeping the feeder layer intact. Although this protocol also works for small clumps, dissociating hESC colonies into single cells allows for a more accurate determination of the number of cells.

Collect dissociated cells 8. Transfer dissociated hESCs in suspension (∼1.5 ml) from the plate to a 15-ml sterile conical tube. Occasionally, small clumps are obtained when dissociating hESC colonies. Dissociate them into single cells using a 1-ml pipet.

9. To recover the maximal amount of dissociated hESCs, wash the dish two times with 1 ml of CMF-DPBS and transfer washes to the same 15-ml sterile conical tube containing dissociated cells.

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10. Centrifuge the tube containing dissociated hESCs in suspension for 5 min at 897 × g, room temperature, to remove all TrypLE Express solution. 11. Remove supernatant and resuspend cells in 500 μl of hESC culture medium containing 8 ng/ml bFGF.

Count cells 12. Count number of viable cells (e.g., using standard Trypan blue staining and a Neubauer chamber or hemacytometer; UNIT 1C.3). Standard size hESC colonies (between 1 and 2 mm in diameter) contain 20,000 cells on average. The total number of cells per plate can be estimated by multiplying the number of colonies dissociated per plate by 20,000.

Plate cells 13. Seed 1500 cells/cm2 onto plates containing fresh feeder layers (see Support Protocol 2 for the preparation of human foreskin fibroblast feeders) and fresh hESC culture medium with bFGF. 14. Add 10 μM ROCK inhibitor Y-27632 stock solution to the culture medium on cells. ROCK inhibitor should be in the culture medium during the first day of culture of dissociated hESCs.

15. Change hESC culture medium with bFGF every day and let the colonies grow for 6 to 8 days.

Prepare dissociated hESCs for freezing 16. After 6 to 8 days, use the stereomicroscope to check the appearance of hESC colonies obtained after expansion of hESCs by single-cell dissociation (Figure 1C.8.4). Almost confluent plates of hESC colonies should be obtained from single-cell splitting and they should present the same features previously described in step 1.

17. If hESC colonies present good morphological features, treat them with 10 μM ROCK inhibitor Y-27632 for 1 hr, and wash and detach them as in steps 2 to 10. 18. After centrifuging the cells, remove supernatant and resuspend pellet in 500 μl of hESC culture medium without bFGF.

A

Cryopreservation of Dissociated hESCs

B

Figure 1C.8.4 Bright-Þeld photographs showing conßuent plates obtained after expansion of hESCs by single-cell dissociation for the cell lines HS207 (A) and HS401 (B). Scale bar = 500 μm.

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19. Count the number of viable cells (e.g., using standard Trypan blue staining and a Neubauer chamber or hemacytometer; UNIT 1C.3). The number of cells obtained after expanding by dissociation of single hESC is two to five times greater (between 8 × 105 and 2 × 106 cells) than the number obtained by mechanical splitting.

Freeze cells 20. Prepare a suspension of at least 100,000 cells/900 μl in cold hESC culture medium without bFGF. Higher amounts of dissociated hESCs can be frozen per cryotube with the same recovery yield.

21. Add 10 μM ROCK inhibitor Y-27632 stock solution to the cell suspension and incubate 10 min at 4◦ C. 22. Prepare 1.8-ml cryotubes by labeling them with hESC cell line, passage number, and date. 23. Add 100 μl DMSO per 900 μl of hESC/culture medium suspension to obtain a final concentration of 10% DMSO. DMSO is commonly used as a cryoprotectant since it protects cells from freezing damage. DMSO is toxic for hESCs at room temperature. All steps involving hESC contact with DMSO at room temperature should be carried out as fast as possible.

24. Mix and transfer 1 ml of cell suspension to each labeled 1.8-ml cryotube. Final cell concentration will be at least 1 × 105 cells/ml of cryopreservation medium.

25. Immediately place cryotubes into a Cryo 1◦ C freezing container and transfer them to a −80◦ C freezer to allow slow freezing (∼1◦ C/min cooling rate). 26. After 24 hr, move tubes to a −150◦ C freezer or liquid nitrogen tank for long-term (years) storage.

Thaw dissociated hESCs 27. Remove a cryovial containing frozen dissociated hESCs from the liquid nitrogen tank or −150◦ C freezer. 28. Rapidly thaw cells by placing cryotube into a 37◦ C water bath. To avoid contamination, the cryotube should not be completely submerged in the water bath, but rather floating with only the bottom of the tube submerged in the water.

29. Agitate cryotube in the bath until almost all the content is thawed. 30. Remove cryotube from the water bath and continue agitating until the cells are completely thawed. 31. Disinfect cryotube by rinsing with 70% ethanol. 32. Immediately transfer thawed cells to a 15-ml sterile tube containing 7 ml hESC culture medium without bFGF. 33. Wash cryotube two times, each time with 1 ml of hESC culture medium without bFGF, to recover all thawed hESCs and transfer them to the same 10-ml sterile tube. 34. Centrifuge thawed cells 5 min at 897 × g to remove all DMSO. 35. Remove supernatant and resuspend cells in 500 μl of hESC culture medium with bFGF.

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36. Count the number of viable cells (e.g., using standard Trypan blue staining and a Neubauer chamber or hemacytometer; UNIT 1C.3).

Plate cells 37. Seed 1500 cells/cm2 onto plates containing fresh mitotically inactivated human foreskin fibroblast feeders in hESCs culture medium containing bFGF. 38. Add 10 μM ROCK inhibitor Y-27632 to culture medium on cells. ROCK inhibitor should be in the culture medium during the first day of culture of cryopreserved dissociated hESCs.

39. On the next day and every day thereafter, change culture medium to hESC culture medium with bFGF and allow colonies to grow for 7 to 10 days. 40. After 7 to 10 days, split hESC colonies mechanically for maintenance culture (see Support Protocol 1). Alternatively, expand them by single-cell dissociation if large numbers of cells are needed to perform experiments (Basic Protocol, steps 1 through 15). SUPPORT PROTOCOL 1

hESC MAINTENANCE CULTURE This protocol describes the preparation of hESC maintenance culture to allow cells to grow for long-term periods while maintaining their properties over time and passages.

Materials hESCs Mitotically inactivated human foreskin fibroblast feeders (see Support Protocol 2) hESC culture medium (see recipe) Scalpels Additional reagents and equipment for hESC culture (see Basic Protocol) 1. Check under a stereomicroscope the appearance of the hESC colonies that are going to be expanded. These colonies should be grown for up to 6 to 8 days on a feeder layer, and they should be of the right size to split. hESC colonies should present the same characteristics described in Basic Protocol, step 1 (Fig. 1C.8.2).

2. If the cells have good morphological features, proceed with mechanical splitting. 3. Cut each colony into 6 to 8 pieces using scalpels and a stereomicroscope (Fig. 1C.8.5). Depending on the size of the hESC colony, the number of pieces obtained per colony will be different. For example, 2-mm diameter hESC colonies should be cut into 8 pieces. Only hESC colonies should be cut into small pieces; avoid cutting into the feeder layer underneath.

4. Repeat step 3 for all the hESC colonies on the plate. If some colonies on the plate have differentiated areas (with undefined borders and are formed by cells with a low nuclear:cytoplasm ratio), do not split them. For an example of differentiated colonies, see Figure 1C.8.3.

Cryopreservation of Dissociated hESCs

5. Collect all the pieces of hESC colonies obtained from one plate with a micropipettor. Seed them onto plates containing fresh feeders in hESC culture medium containing bFGF. It is very important to seed each piece spaced apart from another and distributed homogeneously around the plate. Seed around two to four pieces/cm2 .

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A

B

C

1 DIV

3 DIV

4 DIV

5 DIV

Figure 1C.8.5 Bright-Þeld photographs showing the mechanical splitting process. (A) hESC colony ready to be split mechanically. (B) The same colony has been cut with scalpels in two pieces (arrows). (C) The two pieces of hESC colony 1, 3, 4, and 5 days in vitro (DIV) after seeding growing over a feeder layer. Scale bar = 500 μm.

6. Label plate with hESC line, passage number, and date. 7. Allow pieces of hESC colonies to attach to the feeders for 48 hr. Do not change the culture medium the day after seeding to avoid removing pieces of hESC colonies that are not yet attached.

8. Allow colonies to grow for 6 to 8 days, changing the culture medium every day (Fig. 1C.8.5). 9. After 6 to 8 days, repeat steps 1 to 8 to passage the cells mechanically again or proceed to Basic Protocol for cryopreservation. Alternatively, expand them by single-cell dissociation if large numbers of cells are needed to perform experiments (see Basic Protocol, steps 1 to 15).

PREPARING HUMAN FORESKIN FIBROBLAST FEEDER LAYERS hESCs grow over a feeder layer of human foreskin fibroblasts to retain their undifferentiating properties. Thus, it is necessary to have fresh feeder dishes inactivated mitotically in order to be able to expand hESCs. This protocol describes how to prepare a feeder layer of human foreskin fibroblasts, commercially available from the ATCC. Alternatively, it is also possible to use derived human skin fibroblast lines (see UNIT 1C.7) or mouse embryonic fibroblasts.

SUPPORT PROTOCOL 2

Materials Human foreskin fibroblasts (ATCC # CRL-2429) Culture medium for feeders (see recipe) Additional reagents and equipment for washing and detaching cells (see Basic Protocol)

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Figure 1C.8.6 Phase-contrast photograph showing conßuent human foreskin Þbroblasts feeder cells at 10× magniÞcation.

Passage human foreskin fibroblast cultures 1. Check under an inverted microscope the appearance of the human foreskin fibroblasts grown in culture medium for feeders that are to be expanded. These cells should have been growing between 6 and 8 days and should have a confluent density (Fig. 1C.8.6). Human foreskin fibroblasts grow on standard uncoated 100-mm plates or 75-cm2 flasks in culture medium for feeders. Human foreskin fibroblasts should be subcultured weekly in order to be able to prepare fresh feeder plates for hESC propagation.

2. Remove old medium from the plate with a pipet. 3. Wash three times with 3 to 4 ml of CMF-DPBS. 4. Trypsinize cells by treating them with 2 ml TrypLE Express (for 100-mm plates) for 5 min at 37◦ C. Depending on the size of the plate, the amount of TrypLE Express should be adjusted.

5. After 5 min, check under the inverted microscope to see if cells are detaching from the dish. If the cells are not detached, incubate for an additional 5 min. 6. Once the cells are detached, add 8 ml of culture medium (2 ml of medium per 0.5 ml TrypLE Express) to the dish and dissociate cells with a pipet. Flush the culture medium over the culture plate walls to remove all the remaining adherent cells.

7. Transfer cell suspension to a 15-ml sterile tube. 8. Wash the plate with an additional 2 ml of culture medium for feeders and transfer the wash to the same 15-ml sterile tube. Cryopreservation of Dissociated hESCs

9. Centrifuge cells 5 min at 897 × g, room temperature.

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Count cells 10. Remove the supernatant and resuspend cells in 2 ml of fresh culture medium for feeders. 11. Count the number of viable cells (e.g., using standard Trypan blue staining and a Neubauer chamber or hemacytometer; UNIT 1C.3). Confluent 100-mm plates yield on average 1–2 × 106 cells/plate.

12. Divide the cells obtained into two tubes—one for continued propagation of the fibroblasts and one for feeder layer preparation.

Propagate human foreskin fibroblasts 13. Seed 5 × 105 cells per 100-mm dish in 10 ml culture medium for feeders. 14. Label plate with passage number and date. Human foreskin fibroblast feeders can be used for up to 10 to 15 passages after the initial thawing of the ATCC stock (assuming that ATCC cells are in early passage).

15. Keep the cells growing for 6 to 8 days and passage them when confluent, repeating steps 1 to 14.

Prepare human foreskin fibroblast feeder dishes 16. From freshly dissociated human foreskin fibroblasts (step 12), prepare a cell suspension of 1.5 × 106 cells/ml in culture medium for feeders. 17. Inactivate human foreskin fibroblasts mitotically (e.g., using γ-irradiation, see UNIT 1C.7). Alternatively, mitotically inactivate fibroblasts by mitomycin C treatment (see UNIT 1C.7). Mitotic inactivation of human foreskin fibroblasts is necessary to prevent their expansion into the mixed culture with hESCs.

18. Once they are inactivated by γ-irradiation, seed 3 × 105 cells in 35-mm dishes with 2 ml/dish fresh culture medium for feeders. This cell density ensures confluent mitotically inactivated feeder plates 24 hr after seeding.

19. One day after seeding, use the inverted microscope to check the feeder dishes for the formation of a monolayer of human foreskin fibroblasts. Replace the medium with 2 ml hESC medium not containing bFGF. Culture medium for feeders contains FBS, which is necessary for the regular growth of human foreskin fibroblast. hESCs cannot be in contact with FBS because they start to differentiate. Thus, it is necessary to change the medium of the feeder monolayer to FBS-free hESC culture medium.

20. Use feeders 24 hr after the medium change (they can be used for up to 6 to 8 days— changing the medium every 5 days). Thirty minutes before using mitotically inactivated feeder plates for hESC culture, it is necessary to replace the medium with freshly made hESC medium containing bFGF.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

hESC culture medium Knockout Dulbeco’s modified Eagle’s medium (KO-DMEM; Invitrogen, cat. no. 10829-018) continued

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20% (v/v) Knockout serum replacement (Invitrogen, cat. no. 10828-028) 2 mM L-glutamine (Invitrogen, cat. no. 35050-033) 0.5% (v/v) penicillin-streptomycin (Invitrogen, cat. no. 15140-122) 1% (v/v) MEM non-essential amino acids (Invitrogen, cat. no. 11140-035) 0.5 mM 2-mercaptoethanol (Invitrogen, cat. no. 208665) Store in 50-ml aliquots up to 15 days at 4◦ C Just before use, for complete medium, add: 8 ng/ml basic fibroblast growth factor (bFGF; R&D Systems, cat. no. 234-FSE/CF) Complete hESCs culture medium containing bFGF should be prepared just before using it. A stock solution of 8 μg/ml of bFGF in KO-DMEM should be prepared in 50 μl aliquots and stored up to 3 months at −20◦ C. Avoid repeated freeze/thaw cycles.

Human foreskin fibroblast feeder culture medium Iscove’s Dulbeco’s modified Eagle’s medium (Invitrogen, cat. no. 21980-032) 20% (v/v) freshly thawed and heat-inactivated fetal bovine serum (FBS) 0.5% (v/v) penicillin-streptomycin (Invitrogen, cat. no. 15140-122) Store up to 15 days at 4◦ C COMMENTARY Background Information

Cryopreservation of Dissociated hESCs

The derivation of hESC lines has generated a great interest in using them not only as a source of stem cells for regenerative medicine and cell therapy (Thomson et al., 1998; Trounson and Pera, 2001; Klimanskaya et al., 2008), but also as an invaluable tool for developmental studies (Pannetier and Feil, 2007). However, a significant obstacle for all those applications arises from extremely poor survival rates associated with freezing protocols. Therefore, the establishment of an efficient and reproducible method for cryopreservation of hESCs is essential for the development and widespread use of these cells. Two different approaches have been used for hESC cryopreservation. The most common one is the conventional slow-freezing and rapid-thawing method using dimethylsulfoxide (DMSO) as a cryoprotectant (Grout et al., 1990; Meryman, 2007). This method is based largely on protocols developed for murine embryonic stem cells. Nevertheless, it performs poorly for undifferentiated hESCs, most of which either differentiate or die (Reubinoff et al., 2001; Richards et al., 2004). On the other hand, vitrification of hESCs by the open pulled straw method using high cryoprotectant concentrations together with flashfreezing in liquid nitrogen leads to higher cell survival rates (Reubinoff et al., 2001; Richards et al., 2004; Li et al., 2008). However, vitrified hESC colonies still suffer from high levels of cell death, slow growth rates, and high levels of differentiation. Moreover, this protocol is

tedious to perform manually and is clearly not suitable for handling bulk numbers of hESCs (Heng et al., 2006). In the last few years, several attempts to improve existing cryopreservation methods have been described. These methods include, addition of trehalose or the caspase inhibitor Z-VAD-FMK to the freezing medium, bulk vitrification with cell strainer, adherent cryopreservation, xeno-free cryopreservation, and controlled-rate freezing among others (Ji et al., 2004; Richards et al., 2004; Ware et al., 2005; Wu et al., 2005; Heng et al., 2007; Li et al., 2008). Although all of these methods partially improve the existing ones, the parameters analyzed and the hESC lines used do not allow comparisons among them. Moreover, none are efficient enough to be the clear preferred cryopreservation protocol choice. Taking into account all of these studies, the freezing method that is described here represents a rapid, simple, and reproducible protocol for the cryopreservation of dissociated hESCs.

Critical Parameters and Troubleshooting Work must be done under sterile conditions, all steps involving cell manipulation must be done in a laminar flow hood, and all materials and solutions used for culture should be sterile. Good cell culture practice (GCCP; Coecke et al., 2005) needs to be applied to obtain optimal results, and to minimize the risks associated with handling human materials.

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hESCs grow at 37◦ C and 5% CO2 and are very sensitive to changes in temperature and CO2 levels. Consider growing conditions when cells are removed from the incubator to be checked, passaged, or cryopreserved. Therefore, as a rule, no more than 5 hESC dishes should be processed at the same time to ensure minimal exposure to changes in temperature and CO2 levels. Additionally, the incubator should not be opened for long periods of time to avoid changes in temperature and CO2 levels. Culture of undifferentiated hESCs passaged either by mechanical splitting or by single-cell dissociation presents some critical challenges. Human foreskin fibroblasts used as a feeder layer to grow hESC colonies, should be prepared at least 2 days before seeding hESCs. Thus, human foreskin fibroblasts should be propagated, mitotically inactivated, and seeded at the correct density to obtain confluent plates. It is critical to change the feeder culture medium 24 hr after seeding, and replace it with hESC medium without bFGF. Moreover, feeder cells should have been in hESC medium at least 24 hr before seeding hESCs on them. Human foreskin fibroblasts can support hESC growth up to 15 passages after the ATCC stock is thawed. Another major parameter to ensure undifferentiated growth of hESCs is a daily medium replacement with freshly prepared hESC culture medium containing bFGF. hESC medium and bFGF that has been stored for 30 days at 4◦ C or longer can lose its beneficial properties and therefore the use of a freshly prepared medium is strongly recommended to improve hESC growth. To propagate hESCs, it is necessary to passage only those colonies possessing undifferentiated morphological features, such as welldefined borders and small cells with a high nuclear: cytoplasm ratio (Fig. 1C.8.2). hESC colonies not having such properties should not be passaged. Cryopreservation also has critical steps to ensure high survival rates and efficiency of colony formation with low recovery times. It is essential to start the cryopreservation protocol with healthy, undifferentiated hESC colonies; therefore, avoid the differentiated ones. The first step of expansion by single hESC dissociation is crucial to obtain a large number of cells that can be cryopreserved and stored for longterm periods, ensuring a higher recovery after thawing. Regarding dissociation of hESC, it should be taken into account that hESC possess a high susceptibility to apoptosis upon cellular detachment and dissociation. Therefore,

two considerations are important for hESC dissociation both for passaging and for cryopreserving. Treatment with the ROCK inhibitor Y-27632 1 hr before dissociation is very important to prevent hESC cell death. TrypLE Express treatment should be as short as possible to avoid cell membrane damage. Another important parameter in the cryopreservation protocol is the freezing step. hESCs should be frozen in cold cryopreservation medium containing ROCK inhibitor and 10% DMSO. hESC suspensions should be frozen in cryotubes in a −80◦ C freezer inside a freezing container, ensuring a cooling rate of 1◦ C/min, which has been reported to be critical for hESC survival. For long-term storage, it is also crucial to keep the frozen hESC in a liquid nitrogen tank or a −150◦ C freezer rather than a −80◦ C freezer. The thawing step should be completed as quickly as possible by introducing cryotubes containing frozen hESCs into a 37◦ C water bath and transferring them quickly to a sterile tube containing warm hESC culture medium to dilute the DMSO. Finally, a critical parameter for cell survival and adherence of cryopreserved hESCs to the feeder layer is the addition of the ROCK inhibitor Y-27632 to the culture medium once the cells are seeded (Martin-Iba˜nez et al., 2008).

Anticipated Results In a previous study by Martin-Iba˜nez et al. (2008), the addition of ROCK inhibitor at different steps of the cryopreservation process was tested, concluding that it is necessary to add the inhibitor before dissociation of hESCs, and in the freezing medium and in the postthawing medium. This protocol is an efficient method that allows the freezing/thawing of large amounts of hESCs and ensures high cell survival rates with hardly any differentiation, which in turn gives rise to shorter recovery times. Using this protocol, 50% of hESCs are recovered alive after cryopreservation, which is significantly higher than the survival rates obtained using established methods, which are between 6% and 30%. Moreover, the protocol detailed here presents a high efficiency of colony formation since up to 100 to 200 colonies are obtained from 20,000 cryopreserved hESCs. Other main challenges in the design of an hESC cryopreservation method are the reduction of the differentiation rates and time of recovery, two of the problems encountered when using traditional cryopreservation protocols that include slow freezing/rapid thawing and vitrification (Reubinoff et al., 2001;

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Ji et al., 2004; Richards et al., 2004; Heng et al., 2007). The protocol described in this unit avoids differentiation after cryopreservation of hESC colonies, confirmed by the failure to detect any differentiated colony either by morphological analysis or using Oct4 staining. Furthermore, this method decreases the time of recovery after cryopreservation significantly because confluent plates of undifferentiated colonies from small numbers of dissociated cryopreserved hESCs (3 × 104 to 5 × 105 ) were obtained using this cryopreservation method, a large number of hESC colonies that are ready to be used for several purposes 7 to 10 days after thawing. Standard methods present longer recovery times, since few undifferentiated colonies are obtained after cryopreservation. Thus, using standard methods, it will be necessary to expand the cells to be able to obtain a large amount of hESCs to carry out experiments. In a previous work by Mart´ın-Ib´an˜ ez et al. (2008), it has been demonstrated that after prolonged culture of frozen/thawed dissociated hESCs, the characteristics of pluripotent cells were conserved, including normal karyotype, morphological features, marker expression (SSEA-4, TRA-1-60, TRA-1-81, and Oct-4), and the potential to differentiate into derivatives of all three germ layers after embryoid body formation.

Time Consideration

Cryopreservation of Dissociated hESCs

To estimate the time necessary for cryopreserving dissociated hESCs in the presence of a ROCK inhibitor, the following should be taken into account: feeder dish preparation, hESC expansion by dissociation of single cells, and cryopreservation. The time required for feeder preparation, taking into consideration that human embryonic fibroblasts grow continuously and are passaged weekly, can be estimated from 2 to 5 days depending on the cell density of the feeders at the start of cryopreservation planning. hESC expansion by dissociation of single cells will take 6 to 8 days from the day they are dissociated. This is the time required for hESC colony growth. Once hESCs have expanded, cryopreservation is complete in 1 day, and on the following day, the cells are transferred to a liquid nitrogen tank or a −150◦ C freezer. Thawing of dissociated hESCs is a rapid step that only requires a few minutes. hESC colonies are ready to be used 7 to 10 days after thawing.

Acknowledgements This study was supported by grants from the Swedish Research Council (O.H.; Sweden) and the Ministerio de Educaci´on y Ciencia (SAF2006-04202, J.M.C.; Spain), the Ministerio de Sanidad y Consumo [RETICS (Red de Terapia Celular), J.M.C.; Spain]. R.M-I. is a fellow from ESTOOLS under the European Union’s Sixth Framework Programme.

Literature Cited Coecke, S., Balls, M., Bowe, G., Davis, J., Gstraunthaler, G., Hartung, T., Hay, R., Merten, O.W., Price, A., Schechtman, L., Stacey, G., and Stokes, W. 2005. Guidance on good cell culture practice. A report of the second ECVAM task force on good cell culture practice. Altern. Lab. Anim. 33:261-287. Grout, B., Morris, J., and McLellan, M. 1990. Cryopreservation and the maintenance of cell lines. Trends Biotechnol. 8:293-297. Heng, B.C., Ye, C.P., Liu, H., Toh, W.S., Rufaihah, A.J., Yang, Z., Bay, B.H., Ge, Z., Ouyang, H.W., Lee, E.H., and Cao, T. 2006. Loss of viability during freeze-thaw of intact and adherent human embryonic stem cells with conventional slowcooling protocols is predominantly due to apoptosis rather than cellular necrosis. J. Biomed. Sci. 13:433-445. Heng, B.C., Clement, M.V., and Cao, T. 2007. Caspase inhibitor Z-VAD-FMK enhances the freeze-thaw survival rate of human embryonic stem cells. Biosci. Rep. 27:257-264. Ji, L., de Pablo, J.J., and Palecek, S.P. 2004. Cryopreservation of adherent human embryonic stem cells. Biotechnol. Bioeng. 88:299-312. Klimanskaya, I., Rosenthal, N., and Lanza, R. 2008. Derive and conquer: Sourcing and differentiating stem cells for therapeutic applications. Nat. Rev. Drug Discov. 7:131-142. Li, T., Zhou, C., Liu, C., Mai, Q., and Zhuang, G. 2008. Bulk vitrification of human embryonic stem cells. Hum. Reprod. 23:358-364. Mart´ın-Ib´an˜ ez, R., Unger, C., Str¨omberg, A., Baker, D., Canals, J.M., and Hovatta, O. 2008. Novel cryopreservation method for dissociated human embryonic stem cells in the presence of a ROCK inhibitor. Hum. Reprod. 23:2744-2754. Meryman, H.T. 2007. Cryopreservation of living cells: Principles and practice. Transfusion 47:935-945. Pannetier, M. and Feil, R. 2007. Epigenetic stability of embryonic stem cells and developmental potential. Trends Biotechnol. 25:556-562. Reubinoff, B.E., Pera, M.F., Vajta, G., and Trounson, A.O. 2001. Effective cryopreservation of human embryonic stem cells by the open pulled straw vitrification method. Hum. Reprod. 16:2187-2194. Richards, M., Fong, C.Y., Tan, S., Chan, W.K., and Bongso, A. 2004. An efficient and safe xenofree cryopreservation method for the storage of

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human embryonic stem cells. Stem Cells 22:779789. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Trounson, A. and Pera, M. 2001. Human embryonic stem cells. Fertil. Steril. 76:660-661. Ware, C.B., Nelson, A.M., and Blau, C.A. 2005. Controlled-rate freezing of human ES cells. Biotechniques 38:879-880, 882-883. Wu, C.F., Tsung, H.C., Zhang, W.J., Wang, Y., Lu, J.H., Tang, Z.Y., Kuang, Y.P., Jin, W., Cui, L., Liu, W., and Cao, Y.L. 2005. Improved cryopreservation of human embryonic stem cells with trehalose. Reprod. Biomed. Online 11:733739.

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Authentication and Banking of Human Pluripotent Stem Cells

UNIT 1C.9

Richard Josephson1 and Jonathan Auerbach1 1

GlobalStem, Rockville, Maryland

ABSTRACT Pluripotent human stem cell lines from embryos or reprogrammed adult cells are not all alike. Cell lines differ widely in their propensity for differentiation, their chromosomal integrity and epigenetic state, immunological proÞles, and their availability for research. It is important that all pluripotent cell lines be protected from loss by being properly banked and authenticated, which will also protect current experimental data by enabling its future reproducibility. This unit considers basic guidelines for banking and authentication of pluripotent stem cells that should be easily implementable within any laboratory. Cell Banking is the disciplined preservation of a cell stock in the originally obtained state, as well as stocks representing the baseline state for experimental efforts. Each of these stocks must be authenticated appropriately. Authentication of pluripotent lines veriÞes Þve properties: the unique identity of the line, its sterility or freedom from contaminating microorganisms and pathogens, the integrity and stability of its genome, its expression of typical markers of the stem cell phenotype, and its pluripotency upon differentiation. This unit lists and compares several assays to verify each of these stem cell line properties. Thanks to recent advances in molecular biology and the availability of state-of-the-art assays from service providers, the time and material costs of banking and authentication are not excessive for the C 2009 by John typical research laboratory. Curr. Protoc. Stem Cell Biol. 11:1C.9.1-1C.9.11.  Wiley & Sons, Inc. Keywords: authentication r cell banking r pluripotent stem cell r working cell bank r master cell bank r identity r sterility r stability r undifferentiated phenotype r pluripotency r GMP r GTP

INTRODUCTION Pluripotent stem cells are valued primarily for their capacity to become something else, whether single mature cell types for cell replacement therapy, or in vitro models of congenital disease, or entire animal embryos. Nevertheless, even the most impressive results cannot be conÞrmed if attention is not paid to banking reproducible stem cell reagents. The potency of a human embryonic stem cell (hESC) or induced pluripotent stem cell (iPSC) line can change over time in culture, without noticeable changes in appearance or in marker expression. In addition, hESC lines are not interchangeable; different lines have different propensities to generate particular fates (Chang et al., 2008; Osafune et al., 2008). Therefore, it is critical for every laboratory to protect the properties of its unique lines and the future veriÞability of current research results by observing proper cell banking and authentication practices.

Authentication simply means conÞrming that a cell line is what it is claimed to be. For hESC, this means establishing the unique identity of the line, its sterility or freedom from contaminating microorganisms and pathogens, the integrity and stability of its genome, that it exhibits typical markers of the stem cell phenotype, and its pluripotency upon differentiation. Authentication requires testing these Þve categories of cell line features, using any or all of a variety of assays (see Table 1C.9.1). SpeciÞc protocols for assaying the phenotype and pluripotency of stem cell lines are supplied elsewhere in this volume. Some of the other assays needed to complete authentication are beyond the capabilities of a typical tissue culture laboratory, but are available from reliable service providers at a reasonable cost. Not always taken seriously, cell line authentication has recently been elevated to a higher status in the hierarchy of experimental concerns. In November 2007, the NIH threw

Current Protocols in Stem Cell Biology 1C.9.1-1C.9.11 Published online December 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c09s11 C 2009 John Wiley & Sons, Inc. Copyright 

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some weight behind the issue when it released its “Notice Regarding Authentication of Cultured Cell Lines.” Leaders of diverse institutes encouraged grant reviewers to consider acceptable experimental practices including evolving authentication methods “in order to protect and promote the validity of the science we support” (Bravo and Gottesman, 2007). In addition, some journals are moving closer to asking for, or even requiring, cell authentication information upon publication. Detailed authentication is necessary to deÞne the starting point or baseline condition of stem cells used in research. However, it is of little help to be able to describe the baseline once the cells have irreversibly moved past it. Careful banking will make cells in the baseline condition available for repeat experiments or to share with colleagues. The crucial factors in cell banking for stem cells are to establish a master cell bank (MCB) at the lowest possible passage number, and to return periodically to the baseline condition via a larger working cell bank (WCB). This unit considers basic guidelines for banking and authentication of pluripotent stem cell lines. These should be easily implementable within any stem cell laboratory. We describe the process of cell banking for the individual laboratory, and lay out the authentication requirements at each step. For each of the Þve aspects of authentication listed above, we discuss the risks to pluripotent stem cells and the assays available to negate them. We will examine how advances in molecular biology can lessen the burden of authentication, as can state-of-the-art assays available from service providers. Lastly, this unit will deal brießy with the complex issues around authentication of human pluripotent stem cells for use in the clinic.

Rees et al., 1981), cell cultures are frequently mixed up in the laboratory. A recent largescale effort to Þngerprint 200 donated hESC cultures revealed that 15% of these had been misidentiÞed (Meisner et al., 2008). This is an unacceptably high rate of error, when we consider that investigators’ publications and reputations, not to mention future patients’ lives, are on the line. Fortunately, human cell lines can be unambiguously identiÞed with little effort or expense. By identifying common sequence variations spread throughout the human genome, molecular biology can uniquely identify all individuals save for identical twins. The same techniques can be used to monitor cell line identity. Short Tandem Repeat (STR) analysis is the most common method of DNA Þngerprinting in use today. This technique uses PCR to amplify short interspersed nuclear elements (SINEs) shared among all humans, then quantiÞes the variable numbers of 2 to 4 basepair repeats on each chromatid, typically by capillary electrophoresis for high resolution. Thirteen SINES on different chromosomes are the standard identiÞers for law enforcement worldwide, and this set forms the basis of most STR assay kits. Laboratories that lack the necessary equipment can quickly and cheaply obtain STR proÞles of their lines by sending 10 to 15 μg of isolated DNA (obtainable from ∼3 million cells) to any of several service providers. Results can typically be obtained within several days for a cost of $300 to $450 per sample. The alleles present at commonly assayed STR loci are reported numerically and can be easily compared with databases of ES line identities (such as the International Stem Cell Registry at the University of Massachusetts Medical School, http://www.umassmed.edu/iscr/index.aspx).

IDENTITY

Authentication and Banking of Human Pluripotent Stem Cells

When undertaking an experiment on human embryonic stem cells, it is important to choose the right line. The provenance of the line can affect its availability to a researcher, and also the work’s eligibility for funding. Choosing the wrong pluripotent line can affect the chance of generating a desired cell type (Chang et al., 2008; Osafune et al., 2008). As human pluripotent cells begin to be used therapeutically, cell line identiÞcation will have to be monitored to guarantee the immunological match to the donor/recipient. However, long after the tumor cell line identity crisis caused by the HeLa line (Nelson-

STABILITY The pluripotent cells of the blastocyst do not undergo great expansion in vivo. Placing the inner cell mass into extended culture creates selective pressure on the cells to adapt to foreign conditions (Andrews et al., 2005). Prolonged expansion of hESC frequently leads to genomic and epigenetic abnormalities. Latepassage hESC lines often show chromosomal gains (Draper et al., 2004), ampliÞcations of subchromosomal regions (Maitra et al., 2005), loss of heterozygosity (LOH; Teh et al., 2005; Langdon et al., 2006), alterations of mitochondrial DNA (Maitra et al., 2005), and changes

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in promoter methylation (Maitra et al., 2005; Allegrucci et al., 2007). The most commonly recognized changes are duplications of the short arm of chromosome 12 and the long arm of chromosome 17, which confer a selective advantage in culture (Draper et al., 2004; Imreh et al., 2006). (Interestingly, these two duplications are also implicated in the progression of germ cell tumors; Looijenga and Oosterhuis, 1999). Such chromosomal changes are commonly associated with enzymatic passaging of hESC (Buzzard et al., 2004; Mitalipova et al., 2005). The chromosomal stability of hESC is most frequently monitored by G-band karyotyping. However, karyotyping is expensive and inconvenient enough that most lines are not checked routinely. In addition, microampliÞcations and microdeletions (deÞned as too small to be detectable in a karyotype) frequently occur in hESC lines and are associated with culture adaptation and phenotypic changes (Maitra et al., 2005; Werbowetski-Ogilvie et al., 2009). Fortunately, new methods for molecular cytogenetics now make it possible to routinely monitor for chromosomal ampliÞcations or deletions as small as a few kilobases. As specialist equipment and knowledge is required for these assays, most stem cell laboratories will want to have them performed by a service provider. As little as 1 μg of genomic DNA is required, and the assays are generally not affected by the presence of DNA from mouse feeder cells. The foremost molecular cytogenetic technique is array comparative genomic hybridization (aCGH). In this method, the chromosomes of the cell line to be tested are fragmented and ßuorescently labeled, while a diploid reference line is similarly fragmented and labeled with a second color. Both genomes are hybridized to an arrayed library of cosmids, BAC clones, or oligonucleotides (Solinas-Toldo et al., 1997). A resulting difference in label intensity for any clone indicates a copy number change in the test genome. One limitation to aCGH is that detection of balanced translocations is generally not possible. A modiÞcation of the aCGH method allows the detection of single-nucleotide polymorphisms (SNP; Wang et al., 1998). SNP genotyping provides similar resolution to aCGH for ampliÞcations and deletions, but in addition can detect allelic changes in the genome resulting in loss of heterozygosity (LOH). LOH is a common feature of precancerous tissue (Teh et al., 2005) and of toxicity in ES cells (Donahue et al., 2006). An SNP genotype also

provides an unambiguous identiÞcation of a cell line; however, unlike STR there is no consensus set of SNPs analyzed by all platforms for identity conÞrmation. The cost of array-based cytogenetics is competitive with standard karyotyping, and varies with the size of the microarray. All of the major array manufacturers, including Affymetrix, Agilent Technologies, Illumina, and Roche NimbleGen, offer high-resolution aCGH or SNP arrays for genome-wide detection of copy number variation (CNV). Arrays of 100,000 elements or more are best for CNV detection; the latest arrays pack over 1 million features into every assay. However, the different microarrays cannot be compared solely by the number of loci queried, or by the average spacing between features. Uniform spacing of features increases the functional resolution, which has been deÞned as the size of the smallest single-copy-number change that can be detected in at least 95% of random locations (Coe et al., 2007). In other words, to know how large an ampliÞcation can slip through your net, you need to know the size of the largest holes. Most arrays intended for genome-wide association studies focus on genic regions, leaving large gaps. At least two manufacturers, Affymetrix and Illumina, offer more uniform arrays of ∼300,000 features along with specialized software designed for molecular cytogenetics. The main drawback of these molecular methods is that samples are drawn from many homogenized cells, so the ability to detect aberrant subpopulations must be demonstrated. Typically, depending upon the resolution of the array and the size of the ampliÞcation or deletion, at least 10% to 20% of the population must carry an abnormality before it can be detected by genotyping arrays. Alongside the resolution of arrays, the interpretation of molecular data is continuously improving. For example, aCGH and SNP analyses use normalization methods that assume a diploid background. Therefore, aCGH is unable to recognize complete triploidy or tetraploidy, which are most commonly observed in tumors rather than in stem cells. However, SNP mapping arrays evaluate not only hybridization intensity but also allele ratios at each locus. Thus, SNP panels can identify tetraploidy directly in cases of unbalanced tetrasomy (triplication of one chromosome plus a single copy of the other), or indirectly when subregions of a chromosome are gained or lost, yielding unusual allelic ratios which indicate a nondiploid background (Gardina et al., 2008).

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In addition to cytogenetic changes, hESC lines in extended culture can exhibit alterations of the mitochondrial genome or the methylated DNA epigenome (Maitra et al., 2005; Allegrucci et al., 2007). Both can now be scanned for alterations using recently developed microarrays or sequencing methods. The entire ∼16-kb mitochondrial genome can be checked via resequencing array or direct sequencing. Genome-wide DNA methylation is assayed by a number of methods, based either upon sodium bisulÞte conversion of nonmethylated cytosines to uracil followed by detection of sequence changes, or upon immunoprecipitation of methylated fragments using anti-5-methyl-cytosine antibodies or methylated DNA–binding proteins. The methylated fragments can then be identiÞed by hybridization to aCGH arrays. Monitoring the DNA methylation state is likely to be useful in assessing the completeness of reprogramming in iPSC lines.

STERILITY

Authentication and Banking of Human Pluripotent Stem Cells

Most bacterial and fungal contaminations of tissue culture are discovered by observation. However, it is necessary to include more sensitive tests in the characterization scheme for cell banking. The growth of fungi and mycoplasmas can be very slow, and the use of antibiotics can mask the presence of bacteria in cultures. Thus, a contamination problem may go undiscovered prior to cryopreservation; this can result in the loss of the cell line. Mycoplasmas are usually not observable under the microscope unless very numerous, but even at low abundance they can have many effects on cell behavior and spread easily to clean cultures. There are essentially two types of tests for contaminants in cell culture: direct culture and PCR. In direct culture tests, a vial of cells is thawed and seeded into several types of culture media to detect aerobic bacteria, anaerobic bacteria, and fungi. Mycoplasmas are detected by seeding on nutrient agar plates, which yield a distinctive colony morphology. These tests are extremely sensitive but can take up to 2 weeks for detection of fungi and 3 to 4 weeks for mycoplasmas. Testing by PCR is much faster, requiring only a few hours, and it is very sensitive but the range of detectable species is smaller. Several commercial PCR kits are available that can detect the eight most common species of mycoplasmas in laboratories; some kits detect as many as forty species. PCR tests have also been developed for bacteria in culture and are available as a service

from several providers. Fungal testing by PCR is being developed for clinical samples but is not yet common for cell lines. Testing for bacterial and fungal contamination should be done with one unopened vial of banked cells, rather than from cells in culture. Thus, a positive result unambiguously indicates that contamination occurred prior to freezing, eliminating uncertainty and the expense of retesting additional vials. A concern related to sterility is the possible presence in the cell line of viral pathogens. To protect laboratory personnel, every human cell line should be tested early on (e.g., at the MCB stage) for human viruses such as HIV, HTLV-I and –II, EBV, and so forth. Very sensitive and rapid detection of these pathogens via PCR is available from several service providers, the best of which are certiÞed for clinical laboratory testing. Cell lines that will be transplanted into experimental mice are usually required by laboratory animal protocols to be tested for mouse viral pathogens. This is usually done using the mouse antibody production (MAP) test, a serological screen that produces results in 3 to 8 weeks. Several alternative PCR panels for mouse or rat pathogens are now available that can be performed by a service provider in a few days (Blank et al., 2004).

PHENOTYPE Authentication of phenotype is performed by checking marker expression against a baseline description of the cell line. The phenotype can be veriÞed by antibody detection of pluripotency biomarkers such as POU5f1, SSEA4, etc. (see UNIT 1B.3), or by gene expression analysis (see UNIT 1B.2). Quantitative differences in expression of these markers may be indicative of greater pluripotency in hESCs (Enver et al., 2005; Laslett et al., 2007). To assess quantitative changes, ßow cytometry or gene expression analysis should be performed in parallel with a reference standard such as the embryonal carcinoma line 2102Ep (Josephson et al., 2007). In addition to gene and protein expression, we have included micro RNA (miRNA) expression in the table of phenotype assays (see Table 1C.9.1). Although these small noncoding RNAs are not yet widely recognized as surrogate markers of pluripotency, a number of miRNAs are speciÞc to undifferentiated hESC (Suh et al., 2004; Tang et al., 2006). miRNAs may have important roles in reprogramming and maintaining pluripotency as a link between transcription factors and the epigenetic state (Kashyap et al., 2009).

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Table 1C.9.1 Menu of Selected Assays for Authentication of Pluripotent Stem Cell Linesa

Method

Material required (typical amount)b

Costc (reagent or kit)

Dye exclusion

Trypan blue

Cell suspension

$20/10,000 tests

Live cell metabolism

Fluorescein diacetate/calcein AM

Cells

$50-$200/1000 tests

Mycoplasmas

PCR

Antibiotic-free culture supernatant or frozen cells

$150-$200/25 tests

DNA stain/direct culture

1 vial cells (≥2 × 106 cells)

$175-$342

Bacteria/fungi

Direct culture

1 vial cells (≥2 × 106 cells)

$75-$150

Bacteria

PCR

1 vial cells (≥2 × 106 cells)

$60-$120

Human viral pathogens

PCR

Frozen or cryopreserved cells (≥2 × 106 cells)

$240-$750

Mouse pathogens

MAP test

Frozen or cryopreserved cells (≥2 × 106 cells)

$500-$850

PCR

Frozen or cryopreserved cells (≥2 × 106 cells)

$190-$470

DNA Þngerprint

Short tandem repeat

0.5-2.5 ng DNA

Protein expression

Immunoßuorescence

Fixed cells

Flow cytometry

Dissociated cells

RT-PCR

1-2 μg total RNA

Microarray

1-5 μg cDNA

$200-$575/array

10 ng - 3 μg total RNA

∼$400 kit

Microarray

0.5-5 μg total RNA

$180-$300/array

Karyotype

Dividing cells 1-3 × 25-cm2 ßasks

$250-$850

Spectral karyotype

Dividing cells 1-3 × 25-cm2 ßasks

∼$750

Comparative genomic hybridization (aCGH)

1-2.5 μg DNA

Characteristic Assay

Costc (service)

Viability

Sterility $50-$150

Identity $1500-$2200/100 reactions

$100-$500

Phenotype

Gene expression

miRNA expression qRT-PCR

$750-$995/50 tests $15-$100 $150$1000 plus array

$250-$520

Stability Chromosomes

$300-$650/array

$250-$400 plus array continued

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Table 1C.9.1 Menu of Selected Assays for Authentication of Pluripotent Stem Cell Linesa , contined

Characteristic Assay

Material required (typical amount)b

Costc (reagent or kit)

Costc (service)

Single-nucleotide polymorphism (SNP)

500 ng DNA

$175-$350/array

$215-$700 plus array

Methylation-speciÞc PCR

1 μg bisulÞte-converted DNA

∼$200/50 reactions bisulÞte kit

Whole-genome methylation array

4 μg input DNA + 4 μg immunoprecipitated

∼$500 MeDIP kit $300-$650/array

$275-$450

Sequencing or resequencing array

10-300 ng genomic DNA

∼$200/array

$500$1200

Method

Stability, continued

Epigenome

Mitochondrial DNA Pluripotency Embryoid body

Cell suspension ∼1 × 106 cells

Teratoma

1 vial cryopreserved cells

$1800$3900

a Acronyms: MAP, mouse antibody production; MeDIP, methylated DNA immunoprecipitation; qRT-PCR, quantitative reverse transcriptase polymerase

chain reaction b Check with service providers for exact amount of material needed and how best to extract, preserve, and ship this material. c All prices are in U.S. dollars. The prices are approximate and vary with the breadth or resolution of the speciÞc test. Customers without access to an institutional core facility, or who need validated testing under a manufacturing quality system, can expect higher prices.

Laboratories that do not have the capability to perform ßow cytometry or microarray experiments often have access to an institutional core facility, and some of these offer their services to outsiders for a fee. There are also numerous commercial service providers for microarray analysis and a few for ßow cytometry.

PLURIPOTENCY

Authentication and Banking of Human Pluripotent Stem Cells

This category refers to the functional detection of pluripotency, either by embryoid body (EB) differentiation in vitro or teratoma formation in vivo (see UNIT 1B.4). Embryoid bodies are easily produced in the tissue culture laboratory, but analysis of these for markers of the three embryonic germ layers is less simple. It can be difÞcult to obtain useful immunostaining from whole-mount EBs, while their small size and fragility are a challenge for sectioning. Most researchers turn instead to gene expression analysis, using RT-PCR for germ layer markers (Draper et al., 2007) or microarray analysis. The latter may give more reliable data, since multiple markers for each germ layer are more easily assayed; this can compensate for the expression of some markers in extraembryonic lineages as well.

Teratomas are relatively easy to produce and count in immunocompromised host animals, but the interpretation of immunostained tumor sections usually requires the help of an expert pathologist. Commercial services exist that will not only produce teratomas but provide histological analysis. The dark side of pluripotency is that culture adaptation can create cell lines that remain undifferentiated in the presence of differentiation signals. In the teratoma assay, regions are sometimes found, after many weeks, that still express Oct-4 or other undifferentiated markers. This can be a sign that the transplanted line is karyotypically abnormal (Werbowetski-Ogilvie et al., 2009). The increased self-renewal capacity of aberrant lines can be measured by the dissociation of EBs to seed growth of secondary EBs (Qu and Feng, 1998).

CELL LINE BANKING The process of cell banking is a concern not just for the large hESC distribution centers established by various nations and institutions. Because stem cell lines exhibit changes during passaging, every laboratory needs an in-house banking system to

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derive hES or iPS line

receive hES or iPS line obtain documentation

mycoplasma

expand 5 passages (no antibiotics)

freeze pre-master bank

thaw 2 vials (no antibiotics)

viability

OK? pass

fail

return to source

return to pre-MCB thaw 2 vials (no antibiotics)

sterility

fail

viability OK?

freeze master cell bank

phenotype pass

thaw 1 vial from master cell bank

expand 5 passages (no antibiotics)

compare to documentation

identity

thaw 1 vial from pre-master bank

expand 5 passages (no antibiotics)

sterility

thaw 2 vials (no antibiotics)

cell expansion

sterility viability

return to MCB

identity

fail

phenotype

freeze working cell bank

stability

OK? pass

pluripotency thaw 1 vial from WCB

expand 20 passages

conduct research

publish

stability

Figure 1C.9.1 A flow chart for the process of banking human pluripotent stem cell lines. The typical laboratory should preserve three stocks of each line: the pre-master bank, the master cell bank (MCB), and the working cell bank (WCB). Each of these banks will need to be authenticated for the characteristics listed in the gray boxes. Only the WCB is used to supply experimental work, and experimental cells should be expanded only within a limited number of passages. Banks intended for the wider distribution of cell lines should follow these guidelines, but expand cells from the WCB to preserve a larger distribution bank with the same authentication requirements. Embryonic and Extraembryonic Stem Cells

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ensure that the research results obtained today will be reproducible in the future. The best way to accomplish this is by maintaining well-characterized cryopreserved cells in a working cell bank to supply cells for research. The WCB itself is derived from a smaller, lowpassage master cell bank, which is used only for renewal of the WCB. For a single laboratory, a simple banking scheme consisting of pre-master, MCB, and WCB stocks is sufÞcient. The requirements for hES cell banks intended for distribution of lines to large institutions or entire nations are more numerous. These include the collection of provenance documentation, maintaining a database of characterization information from the originating laboratory, and preserving archival samples (Stacey, 2004). The rules for clinical use of hESC or iPSC lines will be even more stringent and are brießy discussed after this section. The process of banking a pluripotent line for the use of a single laboratory is illustrated in Figure 1C.9.1. The most important consideration is to freeze away a safety stock or pre-master cell bank consisting of Þve to ten vials as soon as possible after obtaining the cell line. The expansion to this level from the initial culture should be done, to the greatest extent possible, following the culture methods recommended by the supplier. Any changes in the protocol or adaptation to new conditions—such as new media or feeders, feeder-free growth, or a new subculturing method—should wait until after the pre-MCB is established and characterized. However, if antibiotics can be eliminated from the medium at this stage it will allow early and accurate sterility testing for bacteria and mycoplasmas. The critical characterization steps for this preMCB are primarily sterility, identity, and viability. Once the pre-MCB is secure, one vial can be expanded 5 passages or less to establish a MCB of 25 to 30 vials. This expansion can be done via the original protocols or ones own. Antibiotics should be omitted from the medium to avoid masking microbiological contamination. It is important to monitor the cells during expansion for signs of change, such as altered growth rates or loss of feeder or mitogen dependence. The characterization done on the MCB should include checks of viability and sterility, along with some assessment of phenotype such as immunocytochemistry. Vials from the MCB are used to expand to a WCB of perhaps 50 to 100 vials, de-

pending upon the number of anticipated users (including colleagues). The WCB is characterized extensively, including measures of sterility, identity, phenotype, stability, and pluripotency. This is done in part because these assays are more expensive not only in terms of money and time, but number of cells required, and it is best to spread these costs over a larger number of vials. Another reason is that the WCB supplies the cells for generating publishable data, so it represents the baseline for changes during handling. In the event that the WCB proves to be karyotypically abnormal or not pluripotent, these assays may then be repeated on the MCB. When experiments require large numbers of cells, these are expanded from the WCB. It is recommended to repeat the characterization of expanded hESC every 10 to 15 passages, especially the stability assays (karyotype or molecular cytogenetics). In addition, although hESC lines have been expanded up to 200 passages, a better method to ensure reproducible cultures is to discard research stocks after 20 to 30 passages and return to the WCB. Finally, the effort of cell banking is wasted if the cell line is preserved but information about it is not. Issues such as provenance and xenobiotic exposure are more likely than biomarker expression levels to restrict the future applications of a line. Also, growing reproducible cultures from a bank may require the same medium, serum, feeders, dissociation methods, etc., as previously used. This information, once lost, cannot be recovered by any assay. Therefore, maintaining complete and accurate records is fundamental to safeguarding a stem cell line.

CLINICAL CONSIDERATIONS Almost from the moment that human embryonic stem cells were Þrst published, researchers in the Þeld have asked regulators for clear guidance on bringing pluripotent cells into the clinic. All cells produced for transplantation must conform to the internationally recognized principles of good tissue practice (GTP) and good manufacturing practice (GMP); however, these principles are embodied differently in the regulations of various nations. Currently, the U.S. Food and Drug Administration (FDA) offers no guidelines speciÞc to hESC lines, regulating them alongside other cultured or extensively manipulated tissues. A thorough review and commentary on the application of FDA rules to the unique issues of pluripotent cells has been recently published (Carpenter et al., 2009).

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The heart of GTP and GMP is maintaining consistent product, which helps to produce predictable outcomes and minimize risks to the patient. This is a great challenge given the heterogeneous and changeable nature of pluripotent stem cells. Under GMP, product consistency is maintained by stringent process control and traceability. Carpenter et al. (2009) propose that hESC derivatives need additional quality control measures beyond those for other cells or tissues. Stem cells being banked or differentiated for cell therapy should be monitored longitudinally for genomic and epigenetic stability, as well as for changes in gene expression and behavior. Animal transplantation studies should be performed on the clinical materials to assess potential tumorigenicity due to residual pluripotent cells or oncogenic transformation. Furthermore, animal or in vitro assays must be developed that are predictive of efÞcacy and potency in vivo. It is worth noting that GMP requires all characterization assays to be validated for accuracy and reproducibility. One question not clearly answered by regulators is where one obtains a hESC line compatible with cell therapy. Crook et al. were the Þrst to publish hESC lines derived specifically for clinical use (Crook et al., 2007). These authors carefully followed the GTP and GMP principles, as well as the FDA requirements for therapeutic cells and international guidelines for ethical procurement of fertilized embryos (see APPENDIX 1A). Their report is essential reading for anyone thinking about deriving or culturing pluripotent stem cells for the clinic. It is required that all culture reagents (media, chemicals, serum or serum substitutes, and even feeder cells) be manufactured under GMP standards and screened for any adventitious agents (bacteria, yeast, mold, mycoplasmas, and extrinsic viruses) that could potentially harm recipients. The authors used human Þbroblasts as feeder cells, but did not forego animal serum because the FDA considers only live animal cells and products from live animal cells to be xenotransplants. The hESC lines were also cultured and banked without the use of antibiotics, which could lead to reactions or resistance problems in patients. Notably, the authors generated over one thousand pages of documentation per hESC line to certify regulatory compliance. Carpenter et al. (2009) point out that complete compliance with current regulations is virtually impossible when deriving new hESC lines. This is because the source materials, leftover in vitro fertilized embryos, are typically

created in fertility clinics rather than GMP manufacturing facilities. However, in at least one case the FDA has allowed cells derived from a research hESC line to enter clinical trials. This approval was made possible by cell banking and rigorous testing for adventitious agents. Carpenter et al. propose that thorough testing should be the requirement instead of GMP-compliant derivation (Carpenter et al., 2009). Finally, the intent of clinical stem cell banking is to minimize variation in the source material. However, to avoid immune rejection in disparate patients, numerous hESC lines or patient-speciÞc iPS lines will have to be differentiated to therapeutic cells. Subtle disparities in pluripotent lines may make it difÞcult to obtain consistent results from standardized differentiation protocols. Therapies based upon iPS technology may bring new sources of variation, such as the age and health of donors, donor cells from different somatic tissues, various methods of reprogramming, and varying completeness of epigenetic reprogramming, as well as the yet-unknown stability of iPS lines. With the current regulatory burden, it seems impractical to create a multitude of clinical-grade patient-speciÞc iPS cell lines to be used once only. This argues that regulators and stem cell scientists should deemphasize the regimentation of processes and push for stronger characterization criteria and rigorous safety and efÞcacy tests. More extensive recommendations for the clinical translation of stem cells have been put forward by the International Society for Stem Cell Research (see APPENDIX 1B).

CONCLUSION Individual practitioners of stem cell culture may feel that cell banking and authentication are expensive indulgences of concern only to repositories. After all, a large part of the art of tissue culture is the portion of every day spent looking down a microscope and observing cells directly. This is often all that is needed to spot emerging problems. Yet, it must be remembered that pluripotent cells undergo invisible adaptive changes to culture conditions, so abnormal cells sometimes grow more robustly and more strongly express stem cell markers. In the absence of standards and standardized guidelines for authentication, ultimately it is the responsibility of each laboratory to decide what authentication assays are necessary for their work and how often to perform these. Many assays too specialized for small tissue culture laboratories are available from service

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providers at a reasonable cost. There is simply no reason to accept the risks of working on cell lines that have not been authenticated. At present, journals have no uniform authentication requirements for publication. However, even published work is vulnerable if the results cannot be reproduced. Reproducing results may require duplicating the exact state of pluripotency in the original reagents, which cannot be done without proper cell banking. The critical factor is to establish a bank of frozen cells as early as possible, to stabilize the line before any changes occur. Complete characterization of the bank establishes a baseline description of the cells, and this will ensure that cells in this known state of authentication are always available. One key difference between stem cells and immortal cell lines is that we intend for stem cells to change over time, in order to generate diverse differentiated progeny. Thus, the quality we are trying to measure and control is not their phenotype, but their potential for good or harm. The products of pluripotent stem cells are just now starting to be used in the clinic, after which the cells may survive and function for the lifetime of the patient. If the Þeld takes time now to establish safe banking and authentication procedures, stem cell medicine may have many good years ahead.

LITERATURE CITED Allegrucci, C., Wu, Y.Z., Thurston, A., Denning, C.N., Priddle, H., Mummery, C.L., Ward-van Oostwaard, D., Andrews, P.W., Stojkovic, M., Smith, N., Parkin, T., Jones, M.E., Warren, G., Yu, L., Brena, R.M., Plass, C., and Young, L.E. 2007. Restriction landmark genome scanning identiÞes culture-induced DNA methylation instability in the human embryonic stem cell epigenome. Hum. Mol. Genet. 16:1253-1268. Andrews, P.W., Matin, M.M., Bahrami, A.R., Damjanov, I., Gokhale, P., and Draper, J.S. 2005. Embryonic stem (ES) cells and embryonal carcinoma (EC) cells: Opposite sides of the same coin. Biochem. Soc. Trans. 33:15261530. Blank, W.A., Henderson, K.S., and White, L.A. 2004. Virus PCR assay panels: An alternative to the mouse antibody production test. Lab. Anim. 33:26-32. Buzzard, J.J., Gough, N.M., Crook, J.M., and Colman, A. 2004. Karyotype of human ES cells during extended culture. Nat. Biotechnol. 22:381-382. Authentication and Banking of Human Pluripotent Stem Cells

Bravo, N.R. and Gottesman, M. 2007. Notice Regarding Authentication of Cultured Cell Lines (Notice Number: NOT-OD-08-017). National Institutes of Health, Bethesda, Md.

Carpenter, M.K., Frey-Vasconcells, J., and Rao, M.S. 2009. Developing safe therapies from human pluripotent stem cells. Nat. Biotechnol. 27:606-613. Chang, K.H., Nelson, A.M., Fields, P.A., Hesson, J.L., Ulyanova, T., Cao, H., Nakamoto, B., Ware, C.B., and Papayannopoulou, T. 2008. Diverse hematopoietic potentials of Þve human embryonic stem cell lines. Exp. Cell Res. 314:29302940. Coe, B.P., Ylstra, B., Carvalho, B., Meijer, G.A., Macaulay, C., and Lam, W.L. 2007. Resolving the resolution of array CGH. Genomics 89:647653. Crook, J.M., Peura, T.T., Kravets, L., Bosman, A.G., Buzzard, J.J., Horne, R., Hentze, H., Dunn, N.R., Zweigerdt, R., Chua, F., Upshall, A., and Colman, A. 2007. The generation of six clinical-grade human embryonic stem cell lines. Cell Stem Cell 1:490-494. Donahue, S.L., Lin, Q., Cao, S., and Ruley, H.E. 2006. Carcinogens induce genome-wide loss of heterozygosity in normal stem cells without persistent chromosomal instability. Proc. Natl. Acad. Sci. U.S.A. 103:11642-11646. Draper, J.S., Smith, K., Gokhale, P., Moore, H.D., Maltby, E., Johnson, J., Meisner, L., Zwaka, T.P., Thomson, J.A., and Andrews, P.W. 2004. Recurrent gain of chromosomes 17q and 12 in cultured human embryonic stem cells. Nat. Biotechnol. 22:53-34. Draper, J.S., S´eguin, C.A., and Andrews, P.W. 2007. Phenotypic analysis of human embryonic stem cells. In Human Embryonic Stem Cells: The Practical Handbook (S. Sullivan, C.A. Cowan, and K. Eggan, eds.) pp. 93-106. Wiley, Chichester, U.K. Enver, T., Soneji, S., Joshi, C., Brown, J., Iborra, F., Orntoft, T., Thykjaer, T., Maltby, E., Smith, K., Dawud, R.A., Jones, M., Matin, M., Gokhale, P., Draper, J., and Andrews, P.W. 2005. Cellular differentiation hierarchies in normal and culture-adapted human embryonic stem cells. Hum. Mol. Genet. 14:3129-3140. Gardina, P.J., Lo, K.C., Lee, W., Cowell, J.K., and Turpaz, Y. 2008. Ploidy status and copy number aberrations in primary glioblastomas deÞned by integrated analysis of allelic ratios, signal ratios and loss of heterozygosity using 500K SNP Mapping Arrays. BMC Genomics 9:489. Imreh, M.P., Gertow, K., Cedervall, J., Unger, C., Holmberg, K., Sz¨oke, K., Cs¨oregh, L., Fried, G., Dilber, S., Blennow, E., and Ahrlund-Richter, L. 2006. In vitro culture conditions favoring selection of chromosomal abnormalities in human ES cells. J. Cell. Biochem. 99:508-516. Josephson, R., Ording, C.J., Liu, Y., Shin, S., Lakshmipathy, U., Toumadje, A., Love, B., Chesnut, J.D., Andrews, P.W., Rao, M.S., and Auerbach, J.M. 2007. QualiÞcation of embryonal carcinoma 2102Ep as a reference for human embryonic stem cell research. Stem Cells 25:437-446.

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Kashyap, V., Rezende, N.C., Scotland, K.B., Shaffer, S.M., Persson, J.L., Gudas, L.J., and Mongan, N.P. 2009. Regulation of stem cell pluripotency and differentiation involves a mutual regulatory circuit of the Nanog, OCT4 and SOX2 pluripotency transcription factors with Polycomb Repressive Complexes and Stem Cell micro-RNAs. Stem Cells Dev. 18:1093-1108 Langdon, J.A., Lamont, J.M., Scott, D.K., Dyer, S., Prebble, E., Bown, N., Grundy, R.G., Ellison, D.W., and Clifford, S.C. 2006. Combined genome-wide allelotyping and copy number analysis identify frequent genetic losses without copy number reduction in medulloblastoma. Genes Chromosomes Cancer 45:47-60.

human embryonic stem cell lines. Nat. Biotechnol. 26:313-315. Qu, C.K. and Feng, G.S. 1998. Shp-2 has a positive regulatory role in ES cell differentiation and proliferation. Oncogene 17:433-439. Solinas-Toldo, S., Lampel, S., Stilgenbauer, S., Nickolenko, J., Benner, A., D¨ohner, H., Cremer, T., and Lichter, P. 1997. Matrix-based comparative genomic hybridization: Biochips to screen for genomic imbalances. Genes Chromosomes Cancer 20:399-407. Stacey, G. 2004. First Report from the UK Stem Cell Bank. National Institute for Biological Standards and Control.

Laslett, A.L., Grimmond, S., Gardiner, B., Stamp, L., Lin, A., Hawes, S.M., Wormald, S., NikolicPaterson, D., Haylock, D., and Pera, M.F. 2007. Transcriptional analysis of early lineage commitment in human embryonic stem cells. BMC Dev. Biol. 7:12.

Suh, M.R., Lee, Y., Kim, J.Y., Kim, S.K., Moon, S.H., Lee, J.Y., Cha, K.Y., Chung, H.M., Yoon, H.S., Moon, S.Y., Kim, V.N., and Kim, K.S. 2004. Human embryonic stem cells express a unique set of microRNAs. Dev. Biol. 270:488498.

Looijenga, L.H. and Oosterhuis, J.W. 1999. Pathogenesis of testicular germ cell tumours. Rev. Reprod. 4:90-100.

Tang, F., Hajkova, P., Barton, S.C., Lao, K., and Surani, M.A. 2006. MicroRNA expression proÞling of single whole embryonic stem cells. Nucleic Acids Res. 34:e9.

Maitra, A., Arking, D.E., Shivapurkar, N., Ikeda, M., Stastny, V., Kassauei, K., Sui, G., Cutler, D.J., Liu, Y., Brimble, S.N., Noaksson, K., Hyllner, J., Schulz, T.C., Zeng, X., Freed, W.J., Crook, J., Abraham, S., Colman, A., Sartipy, P., Matsui, S., Carpenter, M., Gazdar, A.F., Rao, M., and Chakravarti, A. 2005. Genomic alterations in cultured human embryonic stem cells. Nat. Genet. 37:1099-1103. Meisner, L.F., Finger, J.M., Salguero, M.L., and Johnson, J.A. 2008. Karyotype Instability and Cell Line Authentication in Embryonic Stem Cell Cultures. Poster presented at the 6th Annual Meeting of the International Society for Stem Cell Research, Philadelphia. Mitalipova, M.M., Rao, R.R., Hoyer, D.M., Johnson, J.A., Meisner, L.F., Jones, K.L., Dalton, S., and Stice, S.L. 2005. Preserving the genetic integrity of human embryonic stem cells. Nat. Biotechnol. 23:19-20. Nelson-Rees, W.A., Daniels, D.W., and Flandermeyer, R.R. 1981. Cross-contamination of cells in culture. Science 212:446-452. Osafune, K., Caron, L., Borowiak, M., Martinez, R.J., Fitz-Gerald, C.S., Sato, Y., Cowan, C.A., Chien, K.R., and Melton, D.A. 2008. Marked differences in differentiation propensity among

Teh, M.T., Blaydon, D., Chaplin, T., Foot, N.J., Skoulakis, S., Raghavan, M., Harwood, C.A., Proby, C.M., Philpott, M.P., Young, B.D., and Kelsell, D.P. 2005. Genomewide single nucleotide polymorphism microarray mapping in basal cell carcinomas unveils uniparental disomy as a key somatic event. Cancer Res. 65:8597-8603. Wang, D.G., Fan, J.B., Siao, C.J., Berno, A., Young, P., Sapolsky, R., Ghandour, G., Perkins, N., Winchester, E., Spencer, J., Kruglyak, L., Stein, L., Hsie, L., Topaloglou, T., Hubbell, E., Robinson, E., Mittmann, M., Morris, M.S., Shen, N., Kilburn, D., Rioux, J., Nusbaum, C., Rozen, S., Hudson, T.J., Lipshutz, R., Chee, M., and Lander, E.S. 1998. Largescale identiÞcation, mapping, and genotyping of single-nucleotide polymorphisms in the human genome. Science 280:1077-1082. Werbowetski-Ogilvie, T.E., Boss´e, M., Stewart, M., Schnerch, A., Ramos-Mejia, V., Rouleau, A., Wynder, T., Smith, M.J., Dingwall, S., Carter, T., Williams, C., Harris, C., Dolling, J., Wynder, C., Boreham, D., and Bhatia, M. 2009. Characterization of human embryonic stem cells with features of neoplastic progression. Nat. Biotechnol. 27:91-97.

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Clump Passaging and Expansion of Human Embryonic and Induced Pluripotent Stem Cells on Mouse Embryonic Fibroblast Feeder Cells

UNIT 1C.10

Odelya Hartung,1 Hongguang Huo,1 George Q. Daley,1,2,3,4,5,6 and Thorsten M. Schlaeger1,2 1

Stem Cell Program, Children’s Hospital Boston, Boston, Massachusetts Harvard Stem Cell Institute, Cambridge, Massachusetts 3 Stem Cell Transplantation Program, Division of Pediatric Hematology Oncology, Children’s Hospital Boston, and Dana-Farber Cancer Institute; Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, Massachusetts 4 Division of Hematology, Brigham and Women’s Hospital, Boston, Massachusetts 5 Howard Hughes Medical Institute, Boston, Massachusetts 6 Manton Center for Orphan Disease Research, Boston, Massachusetts 2

ABSTRACT The ability of human embryonic stem cells (hESCs) to differentiate into essentially all somatic cell types has made them a valuable tool for studying human development and has positioned them for broad applications in toxicology, regenerative medicine, and drug discovery. This unit describes a protocol for the large-scale expansion and maintenance of hESCs in vitro. hESC cultures must maintain a balance between the cellular states of pluripotency and differentiation; thus, researchers must use care when growing these technically demanding cells. The culture system is based largely on the use of a proprietary serum-replacement product and basic fibroblast growth factor (bFGF), with mouse embryonic fibroblasts as a feeder layer. These conditions provide the basis for relatively inexpensive maintenance and expansion of hESCs, as well as their engineered counterparts, human induced pluripotent stem cells (hiPSCs). Curr. Protoc. Stem Cell C 2010 by John Wiley & Sons, Inc. Biol. 14:1C.10.1-1C.10.15.  Keywords: human embryonic stem cells r human induced pluripotent stem cells r mouse embryonic fibroblasts r bFGF

INTRODUCTION This unit describes a protocol for the abundant expansion of karyotypically stable human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs) on primary mouse embryonic fibroblasts (MEFs). hESCs were originally derived in medium containing 20% fetal bovine serum, on tissue culture plates coated with inactivated MEFs (Thomson et al., 1998). Current hESC medium recipes substitute serum with a proprietary serum-replacement and supplement this medium with bFGF. hESCs are regularly passaged by mechanical- and enzyme-based methods. Both methods require that the cultures be passaged as small clumps of cells in order to mitigate dissociation-induced cell death and to maintain chromosome stability. This unit will describe both mechanical (Basic Protocol 2) and enzymatic (Basic Protocol 1) passaging methods, as well as methods for thawing (Support Protocol 1) and cryogenically preserving (Support Protocol 2) hESCs. The protocols and procedures are also applicable to human induced pluripotent stem cells (hiPSCs), which closely resemble hESCs in their behavior, epigenetic state, and developmental potential.

Current Protocols in Stem Cell Biology 1C.10.1-1C.10.15 Published online August 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c10s14 C 2010 John Wiley & Sons, Inc. Copyright 

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NOTE: All protocols in this unit require standard tissue culture and sterilization facilities. Cells should be handled under sterile conditions in a Class II Biological biosafety cabinet. NOTE: All human and mouse-derived cells are incubated at 37◦ C in a humidified atmosphere of 5% CO2 in air. NOTE: All equipment and reagents that come into contact with live cells must be sterile, and proper aseptic technique should be used. BASIC PROTOCOL 1

BULK PASSAGING AND EXPANSION OF HUMAN EMBRYONIC STEM CELLS (PICK TO REMOVE) hESCs can be cultured indefinitely in vitro, but there is evidence to suggest that over time, enzymatic passaging of hESCs may promote the emergence of karyotypically abnormal cells, in particular when colonies are dissociated into single cells (Draper et al., 2004). Ideally, the cells can be perpetually maintained by manual scoring and dissection of the undifferentiated hESC colonies, followed by replating of the clumps onto fresh MEF-coated tissue culture dishes (Basic Protocol 2). However, this passaging method is time-consuming and tedious, and can be impractical for the large-scale expansion of cells necessary for experimentation. Alternatively, enzyme-facilitated passaging methods that maintain the cells in small aggregates reduce the cellular stresses that cause chromosomal instability. These methods allow for the bulk passaging of hESCs, and have been used routinely by the authors on various stem cell lines for over 30 passages with no apparent consequences for karyotype, differentiation capacity, or molecular signatures. When using either mechanical or bulk passaging methods, the time between passaging is critical for the overall health of the culture (see Troubleshooting). Robustly growing hESC lines should be regularly passaged approximately every ∼5 to 7 days at split ratios of ∼1:3-1:8. Different hESC lines may have slightly different morphology and growth rates, and may therefore have variable optimal passaging periods and ratios. All hESCs should have their medium changed daily to accommodate their high metabolism and to replenish unstable medium components.

Materials 0.1% (w/v) gelatin solution (Millipore, cat. no. ES-006-B) iMEF medium (see recipe) 70% (v/v) isopropanol in water (VWR, cat. no. EM-PX1834-1) Mitotically inactivated (gamma irradiated) CF-1 MEFs (iMEFS; Global Stem, cat. no. GSC-6001G), frozen Confluent tissue culture well (9.6 cm2 or greater) of hESCs on MEFs hESC medium (see recipe) DMEM/F-12 (Stem Cell Technologies, cat. no. 36254) 1 mg/ml collagenase IV (see recipe)

Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

6-well tissue culture plates (VWR, cat. no. 73520-906) 37◦ C incubator 15- and 50-ml conical centrifuge tubes (BD Biosciences, cat. nos. 352097 and 352070) Tube rack Centrifuge Stereomicroscope (e.g., Discovery V8 with transmitted light darkfield base, Zeiss) Vacuum aspirator Cell lifter (Corning, 3008) 5-ml single-use glass serological pipets (VWR, cat. no. 93000-696) 2-, 5-, 10-, 25-ml plastic serological pipets (VWR, cat. no. 53283)

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Prepare iMEF-coated tissue culture plates 1. Prepare tissue culture plates by dispensing 0.1% gelatin solution onto the plating surface at a volume of 1 ml/10 cm2 (1 ml/well of 6-well plate). Coat surface entirely, and allow plates to incubate for at least 20 min at 37◦ C. 2. Prepare a 50-ml centrifuge tube with 10 ml of iMEF medium and another 50-ml centrifuge tube with 45 ml of 70% isopropanol in water. Tightly close and prewarm both tubes by placing them upright into a tube rack in a tissue culture incubator for at least 20 min. 3. Thaw frozen iMEFs by immersing the bottom of the cryovial in the prewarmed 70% isopropanol until its contents have almost completely thawed (occasional gentle flicking of the cryovial is recommended) and a very small piece of ice remains frozen. The authors prefer using prewarmed isopropanol, as opposed to a water bath, to ensure culture sterility. The preferred mouse strain for the preparation of mouse feeder layers for the support of hESCs is CF-1. CF-1 iMEF-coated plates should be prepared the day prior to use, but can be prepared a few days in advance. Requisite iMEF densities vary greatly from batch-to-batch; each batch of iMEFs should be tested separately for optimal density (see Critical Parameters and Troubleshooting). Always wash iMEFs once with DPBS or DMEM/F-12 before plating hESCs in order to remove iMEF medium components.

4. Wipe off the outside of the tube, remove the lid, and quickly add 500 μl of the prewarmed iMEF medium in a dropwise manner. Aspirate and transfer the entire volume to 50 ml of iMEF medium. 5. Centrifuge the cells 4 min at 200 × g, room temperature. Aspirate the supernatant, and carefully and completely resuspend cells in desired volume of iMEF medium (∼2 ml per 200,000 iMEF cells). 6. Completely aspirate the gelatin solution from the wells, and then distribute the suspended iMEFs according to the batch-dependent optimized density (∼2 ml or 200,000 cells per 10 cm2 ). Transfer iMEF plates to the incubator.

Remove differentiated hESCs These procedures are performed under a stereomicroscope, in a sterile tissue culture biosafety cabinet. It may be necessary to modify the sash of the biosafety cabinet to allow access to the microscope. 7. Feed the confluent hESC culture wells with fresh, prewarmed hESC medium (2 ml/6 well) 1 to 2 hr prior to passaging. 8. Observe the culture under the stereomicroscope for areas of differentiation (refer to Fig. 1C.10.1A-D). 9. Using a vacuum aspirator operating at low speed, carefully aspirate differentiated cells, replenishing with prewarmed DMEM/F-12 as medium is depleted to avoid having the hESCs dry out (Fig. 1C.10.2A-C). For vacuum aspiration (Fig. 1C.10.3), connect a T-connector between the aspirator tubing and a valve that leads to a 1000-μl filter tip (for fine control of vacuum speed). Attach a filter-free 1000-μl tip (e.g., Rainin, cat. no. RT-L1200) to the end of the tubing; this tip can then be fitted with a sterile 12.5-μl pipet tip (e.g., Matrix, cat. no. 7422) for direct aspiration. The ideal setting produces a low but continuous (no back-flow) aspiration speed of less than 50 μl per sec.

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A

B

C

D

Figure 1C.10.1 Morphology of hESCs cultured on iMEFs. An hESC culture in good condition will display the standard hallmarks of hESC colonies, including defined borders and homogeneous cell types within the colonies. (A) hESC culture at optimal passage point. It is normal for hESC cultures to have up to 10% spontaneous differentiation. Types of spontaneous differentiation vary, but they include central differentiation (B, C) and peripheral differentiation (D). Scale bar = 250 μm.

Passage the cells 10. Wash the cells once with 1 ml prewarmed DMEM/F-12, and then add 1 ml/10-cm2 of 1 mg/ml collagenase IV splitting solution. 11. Incubate 3 to 15 min at 37◦ C, until the edges of colonies are visibly curling (Fig. 1C.10.4), the iMEFs are visibly detaching, or the 15-min mark has been reached. Skip step 12 if colonies have already detached completely.

12. Optional: Aspirate the enzyme, and carefully add 1 ml/10-cm2 DMEM/F-12 without detaching the now loosely adherent colonies. Use the cell lifter to gently scrape off the colonies in large clumps. 13. Collect the clumps using a single-use 5-ml serological glass pipet, and transfer the fragments to a 15-ml conical tube.

Wash the cells 14. Wash the well with 1 ml DMEM/F-12 to collect additional fragments and transfer them to the tube (from step 13). Avoid unnecessary pipetting to minimize trituration at this step. Centrifuge the fragments 1.5 min at 200 × g, room temperature. 15. Aspirate the supernatant and resuspend the cells in a suitable volume of hESC medium to achieve a suitable split ratio. Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

hESCs are typically split at ratios ranging from 1:3-1:8 (ratios based on growth area). For example, a single 6-well (∼10 cm2 ) should be resuspended in a total volume of 12 ml hESC medium to achieve a split ratio of 1:6 (2 ml per well).

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Figure 1C.10.2 Demonstration of pick-to-remove (A-C) and pick-to-keep (D-F) methods. For pick-to-remove, remove areas of spontaneous differentiation (A) using a slow vacuum aspirator attached to a filter-free 12.5-μl pipet tip (B,C). Aspirating the differentiation area slowly avoids excessive medium depletion and removal or damage to undifferentiated hESCs. For mechanical passaging, use the tip of a 27-G needle to score undifferentiated colonies into small fragments (E). Once colonies have been scored, carefully push fragments off and collect them using a 200-μl tip (F). Scale bar = 500 μm.

(air) 1000- l tip (filter)

valve

12.5- l tip (no filter)

Figure 1C.10.3

1000- l tip (no filter)

T-connector

(to waste bottle,vacuum)

Diagram of a slow-vacuum aspirator.

16. Triturate cells to break down the clumps until they contain on average about 100 to 200 cells. Clumps of this size are just large enough to be visible in your pipet. Optimal trituration requires a pipet bore size that is not too small and not too large. 5- to 10-ml serological pipets as well as 2000-μl pipet tips work best. It is important that the pipet be able to hold the entire volume of medium that is present during trituration, to allow the entire suspension to be pipetted up and down (a small volume should remain in the tube to avoid formation of bubbles). If necessary, perform trituration in a smaller volume followed by addition of medium to the desired volume. Cell clumps that have been over-triturated will result in reduced plating efficiencies. Conversely, producing clumps that are too large will inhibit expansion and lead to cultures with excessive differentiation, as well as formation of colonies of unequal sizes that may not grow synchronously, and which therefore may not be ready to be passaged simultaneously.

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Figure 1C.10.4 A colony of hESCs that has been incubated in collagenase IV splitting solution for 15 min. Scale bar = 250 μm.

If the hESC colonies were inadvertently triturated down to very small clumps or even single cells, it is advisable to add the Rho-Kinase (ROCK) inhibitor Y-27632 (Sigma, cat. no. Y0503) to a final concentration of 5 μM during plating in order to limit dissociation-induced apoptosis. However, the use of Y-27632 should be limited, as the long-term effects on hESCs have not been fully characterized.

Plate the cells 17. Before plating hESCs onto iMEFs, wash iMEF wells once with 1 ml DMEM/F-12. Add 1 ml of hESC medium to each well. 18. Plate the triturated cells one well (2 ml) at a time, mixing the suspension in between, to achieve an even number of cells across parallel wells. Taking up the entire suspension all at once will likely result in uneven plating between wells since the larger cell clumps settle quickly in the pipet. It is recommended to add the cell clumps dropwise throughout each well. Avoid swirling of the medium to prevent clumps from accumulating in the well center.

19. Transfer the plate to the incubator, and rock the plate from left to right, then back and forth (never in a circular motion), to distribute the cells evenly across the plating surface. Change the medium daily and passage cells every 5 to 7 days once the cultures reach near-confluency (judged as ∼70% growth area). BASIC PROTOCOL 2

Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

MECHANICAL PASSAGING OF HUMAN EMBRYONIC STEM CELLS (PICK TO KEEP) Mechanical passaging, by manual dissection of undifferentiated hESC colonies, is a commonly used method for propagating hESC cultures. Mechanical passaging is the preferred method when more than 20% to 30% of the colonies show signs of differentiation—at which point it becomes impractical to remove all differentiation by vacuum aspiration. This method is therefore particularly useful for recovering the overall “health” of a culture (as judged by percent differentiation). Mechanical passaging is also the method of choice when only a small number of colonies are present, as may be the case after a thaw.

1C.10.6 Supplement 14

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Materials Plate or well of hESCs on iMEFs hESC medium (see recipe) DMEM/F-12 (Stem Cell Technologies, cat. no. 36254) Stereomicroscope (e.g., Discovery V8 with transmitted light darkfield base, Zeiss) 27-G needle, 1/2 -in. long (BD, cat. no. 305109) Syringe, 1-ml or 3-ml volume (BD, cat. nos. 309628 and 309585) Pipet Lite LTS Pipettor, 20 to 200 μl volume (Rainin, cat. no. L-200) Pipet Lite 200-μl filter tips (Rainin, cat. no. RT-L200F) 15- and 50-ml conical tubes (BD Bioscience, cat. nos. 352097 and 352070) 5-, 10-, or 25-ml plastic serological pipets (VWR, cat. no. 53283) Additional reagents and equipment for preparing iMEF-coated tissue culture plates (Basic Protocol 1) 1. Feed the well(s) to be passaged with fresh prewarmed hESC medium (2 ml/6-well) 1 to 2 hr before passaging. If there are wells that are not ready to be passaged, they should be fed as normal until they are ready.

2. Observe the culture under a stereomicroscope, and select 15 to 20 colonies per well that have maintained pristine hESC morphology or that at least contain extended regions without overt differentiation. These procedures are performed under a stereomicroscope, in a sterile tissue culture biosafety cabinet.

3. Using a 27-G needle that has been attached to a 1-ml syringe and bent at a 45◦ angle, carefully cut all undifferentiated areas into a grid containing approximately 9 (3 × 3 grid) to 25 (5 × 5 grid) squares per medium-sized colony (Fig. 1C.10. 2D,E). Be sure to exclude any areas of the colony that are overtly differentiated. Clumps should be ∼100 to 200 cells/clump.

4. Using a 200-μl pipet, slowly dislocate each square (by pushing/scraping with the end of the tip) and collect the clumps (Fig. 1C.10.2F). Dispense the clumps into a 15-ml conical tube containing a few milliliters (2 to 3 ml) prewarmed DMEM/F12 without breaking them up any further. 5. Once all the clumps from the passaged cell line (1 or more wells) have been collected, centrifuge the tube 1.5 min at 200 × g, room temperature. 6. Meanwhile, prepare fresh iMEF plates by aspirating the iMEF medium and washing each well once with 1 ml prewarmed DMEM/F-12 (see Basic Protocol 1 steps 1 to 6 for the preparation of iMEFs). Dispense 1 ml of hESC medium into washed wells. 7. Aspirate the supernatant above the centrifuged hESC pellet, and add fresh prewarmed hESC medium (1 ml per 10 cm2 or 6-well plate well to be plated), and gently resuspend the pellet. No trituration is necessary or recommended here, since the clumps will already be at the appropriate size for replating. The optimal resuspension volume depends on the desired split ratio, which in turn depends on the quality and quantity of the mechanically picked colony fragments. Most undifferentiated colony fragments should be able to attach and initiate the growth of a new, undifferentiated colony. About 100 to 300 fragments should be plated into a 10-cm2 well of a 6-well plate.

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8. Plate the clumps, one well (2 ml) at a time, carefully mixing the suspension in between wells to ensure even plating across all wells. Taking up the entire suspension all at once will likely result in uneven plating between wells since the larger cell clumps settle quickly. It is recommended to add the cell clumps dropwise throughout each well. Avoid swirling of the media to prevent clumps from accumulating in the center of the well.

9. Transfer the plate to the incubator, and rock the plate from left to right, then back and forth, to distribute the cells evenly across the plating surface. Change the medium daily and passage cells every 5 to 7 days once the cultures reach near-confluency (judged as ∼70% growth area). SUPPORT PROTOCOL 1

FREEZING STOCKS OF HUMAN PLURIPOTENT STEM CELLS hESCs can be cryopreserved in a liquid nitrogen freezer for long-term storage. In this protocol, cells are suspended in a DMSO-based cryoprotectant medium, and then frozen by slow cooling to −80◦ C at a rate of ∼1◦ C/min in a −80◦ C freezer, followed by transfer to the gas phase of a liquid nitrogen freezer. hESCs are generally frozen at a ratio of one confluent well of a 6-well plate per cryovial, approximately 1 to 2 million cells per vial.

Materials 2× hESC freezing medium (see recipe) Confluent tissue culture well (9.6 cm2 or greater) of hESCs on iMEFs hESC medium (see recipe) Mr. Frosty (Nalgene, cat. no. 5100-0001) Cryovials (Nalgene, cat. no. 5000-1020) −80◦ C freezer Liquid nitrogen cryostorage unit Additional reagents and equipment for collecting hESCs (Basic Protocol 1) 1. Prepare 250 μl of 2× freezing medium per well to be frozen. Chill 2× freezing medium and Mr. Frosty on ice for at least 30 min prior to use. The authors have tested the addition of 10 μM of ROCK inhibitor Y-27632 to the freezing medium and have not found improved recovery.

2. Collect hESC as described in Basic Protocol 1, steps 7 to 14. 3. After centrifugation, aspirate the supernatant and carefully resuspend cell clumps in 250 μl cold hESC medium per one well of a 6-well plate (or ∼10 cm2 ) of confluent hESCs collected. Avoid any further trituration (usually the cell clumps can be brought into suspension by flicking the bottom of the tube; if pipetting is necessary, use a wide bore tip, such as a 2000-μl pipet). 4. Working quickly, add 250 μl of cold 2× hESC freezing medium per one 6-well to the cell suspension, and mix gently by flicking the bottom of the tube. The final concentration of DMSO is 10%.

5. Dispense 500 μl of the cell mixture per prelabeled cryovial, and place vials in a chilled Mr. Frosty. Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

6. Transfer the Mr. Frosty containing the cryovials to a −80◦ C freezer for 24 to 48 hr for slow freezing, then transfer vials to the gas phase of a liquid nitrogen cryostorage unit.

1C.10.8 Supplement 14

Current Protocols in Stem Cell Biology

THAWING CRYOGENICALLY PRESERVED HUMAN EMBRYONIC STEM CELLS

SUPPORT PROTOCOL 2

hESCs are thawed by rapidly bringing the cells to 37◦ C and removing residual cryoprotectants. Unfortunately, the survival rate of hESCs following such freeze/thaw methods remains inefficient and inconsistent between cell lines and batches. Depending on the length of time frozen, and condition of hESCs at the time of freezing, it may take a few days to several weeks to see the emergence of hESC colonies after thawing.

Materials hESC medium (see recipe) 70% (v/v) isopropanol in water Frozen hESCs (Support Protocol 1) 6-well tissue culture plates of iMEFs (see Basic Protocol 1) DMEM/F-12 (Stem Cell Technologies, cat. no. 36254) 15- and 50-ml conical tubes (BD, cat. nos. 352097 and 352070) Centrifuge NOTE: The authors have found that the addition of 10 μM of ROCK inhibitor Y-27632 to the plating medium might greatly increase cell recovery after a thaw (see UNIT 1C.8). 1. Prepare two separate aliquots of hESC medium (3 and 9 ml) in 15-ml conical tubes, as well as another 50-ml conical tube with 45 ml of 70% isopropanol in water. Tightly close and prewarm the tubes by placing them in a tissue culture incubator for at least 20 min. 2. Quickly thaw the frozen hESCs by immersing the bottom of the cryovial in the prewarmed 70% isopropanol until its contents have almost completely thawed (occasional gentle flicking of the cryovial is recommended) and a very small piece of ice remains frozen. 3. Wipe off the outside of the tube and quickly add 500 μl of the prewarmed hESC medium (from 9-ml aliquot) in a dropwise manner into the tube. 4. Collect the entire contents of the tube into the remaining medium (avoid triturating the cell clumps any further). Centrifuge the cell suspension 1.5 min at 200 × g, room temperature. 5. Meanwhile, wash one well of iMEFs with DMEM/F-12 and add 1 ml of hESC medium from the tube containing the 3-ml aliquot. 6. Aspirate supernatant and gently resuspend pellet using the remaining 2 ml of prewarmed hESC medium from the tube containing the 3-ml aliquot, taking care not to triturate the hESC clumps any further. 7. Plate the cells in the prepared iMEF well. Transfer the plate to the incubator, and shake the plate from left to right, then back and forth (avoid circular swirling), to distribute the cells evenly across the plating surface. 8. Completely change the medium to fresh prewarmed hESC medium 36 to 48 hr after thawing and daily thereafter. Expect colonies to appear within 7 to 14 days. 9. After thawing, if there are only a few undifferentiated high-quality hESC colonies, do the first passaging mechanically and at a low split ratio (it may even be necessary to passage the colonies onto a smaller well). Enzymatic clump passaging may resume as soon as the culture has recovered and begun to grow robustly without excessive differentiation.

Embryonic and Extraembryonic Stem Cells

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Supplement 14

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. Be sure to exclusively use cell culture–grade reagents and consumables—hESCs are exquisitely sensitive to contaminants. Consistent success with hESC culture requires rigorous testing of any new batch of reagents, in particular KOSR, FBS, iMEFs, and collagenase IV. We generally avoid using antibiotics for hESC culture. For suppliers, see SUPPLIERS APPENDIX.

Gelatinized culture plates Prior to plating iMEFs, coat all plates with enough 0.1% gelatin solution (Millipore, cat. no. ES-006-B) to cover the surface. Incubate the plates for at least 20 min in 37◦ C. Completely remove gelatin solution before using.

Human basic fibroblast growth factor (bFGF) stock (10 μg/ml, 1000×) Reconstitute 10 μg bFGF (Gemini Bio-Products, cat. no. 400-432P) in 1 ml diluent solution (see recipe). Use immediately or freeze in 500-μl aliquots for up to 3 months at −20◦ C.

Collagenase IV stock (10 mg/ml, 10×) Dissolve 1 g of collagenase IV (Invitrogen cat. no. 17104-019) in 100 ml DMEM/F12 medium (Stem Cell Technologies, cat. no. 36254)). Mix until dissolved. Filtersterilize the resulting 10× medium and freeze in 1.5-ml aliquots for up to 6 month at −20◦ C. The final concentration of collagenase splitting solution is 1 mg/ml: add 13.5 ml DMEM/F-12 medium to 1.5 ml collagenase IV stock. Mix well. Use immediately or store up to 2 weeks in 4◦ C. Warm aliquot to 37◦ C prior to use.

Diluent solution (0.1% w/v BSA in CMF-DPBS) Add 134 μl of 7.5% (w/v) bovine serum albumin (BSA; Invitrogen, cat. no. 15260) in 10 ml DPBS without Ca2+ or Mg2+ (CMF-DPBS; Invitrogen, cat. no. 14190). Filter-sterilize the resulting solution and use immediately.

Heat-inactivated fetal bovine serum (FBS) Put the frozen bottle of FBS (Gemini Bio-Products, cat. no. 100106) into a 37◦ C water bath until completely thawed (occasionally shake the bottle to facilitate thawing). Fill an empty bottle of equal size with cold (4◦ C) water and insert a thermometer. Set the water bath temperature to 55◦ C and incubate both bottles for ∼30 min, counted from the time the water bottle first reaches ∼55◦ C (occasionally shake both bottles to ensure even heating). Transfer the heat-inactivated FBS bottle to a bucket with water and ice. Store cooled bottle at 4◦ C; use for up to 6 months.

Irradiated mouse embryo fibroblast (iMEF) medium DMEM containing: 10% heat-inactivated FBS (v/v; see recipe) 2 mM L-glutamine (Invitrogen, cat. no. 25030-081) 1 mM sodium pyruvate (Invitrogen, cat. no. 11360-070) 100 μM MEM-nonessential amino acids (Invitrogen, cat. no. 11140-050) 50 U/ml penicillin, 50 μg/ml streptomycin (Invitrogen, cat. no. 15140), optional Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

To prepare 500 ml of the medium, mix 50 ml of heat-inactivated FBS, 5 ml of L-glutamine, 5 ml of sodium pyruvate, 5 ml of MEM- nonessential amino acids, and 5 ml of penicillin/streptomycin (optional), and then fill to 500 ml with DMEM (Invitrogen, cat. no. 11965-092). Protect from light. Store up to 1 week in 4◦ C.

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Current Protocols in Stem Cell Biology

hESC medium DMEM/F12 containing: 20% (v/v) KnockOut Serum Replacement (KOSR; Invitrogen, cat. no. 10828028)10 ng/ml bFGF (see recipe) 1 mM L-glutamine (Invitrogen, cat. no. 25030-081) 100 μM MEM-nonessential amino acids (Invitrogen, cat. no. 11140-050) 100 μM 2-mercaptoethanol (Sigma, cat. no. M7522) 50 U/ml penicillin, 50 μg/ml streptomycin (Invitrogen, cat. no. 15140), optional To prepare 500 ml of the medium, mix 100 ml of KOSR, 5.0 ml of L-glutamine, 5 ml of MEM-nonessential amino acids, 3.5 μl of 2-mercaptoethanol, 5 ml of penicillin/streptomycin (optional), 500 μl bFGF stock (see recipe), and then fill to 500 ml with DMEM/F12 (StemCell Technologies, cat. no. 36254). Protect from light. Store up to 1 week in 4◦ C.

hESC freezing medium, 2× Defined FBS (Hyclone, cat. no. SH30070.01) containing 20% dimethyl sulfoxide (DMSO; Sigma, cat. no. D2650). Chill on ice prior to use. Store up to 1 week in 4◦ C.

COMMENTARY Background Information Understanding the exogenous cellsignaling components that regulate hESC pluripotency is an active field of hESC research. In addition to the standard KOSR/MEF -based culturing conditions, other chemically defined and feeder-free culture conditions have been reported to sustain hESC pluripotency (e.g., Liu et al., 2006; Ludwig et al., 2006). Although these conditions vary, common themes among them include the use of bFGF, insulin/IGF, and activin/nodal/TGFβpathway agonists, as well as lack (or inhibition) of BMP. In KOSR/MEF-based culturing conditions, bFGF together with factors either contained in KOSR (a proprietary product that contains insulin, amino acids, trace elements, anti-oxidants, albumin, lipids, and transferrin), or produced by iMEFs, provide a supportive environment for expansion of undifferentiated hESCs. The authors have forged very distinct roles for mechanical passaging and enzymatic passaging throughout the maintenance of hESC or hiPSC cell lines. The first passage after the cells have thawed is always done mechanically, as the cells will react adversely to enzyme treatment at this time (with increased differentiation and reduced plating). After the first one or two mechanical passages, all cell lines are maintained with enzymatic passaging. A high degree of differentiation or reduced plating may also warrant a couple of mechanical passages during maintenance.

The authors routinely use chemically defined feeder-free culturing conditions (mTeSR1/Matrigel; Ludwig and Thomson, 2009) and KOSR/MEF-based culturing conditions for their daily maintenance and expansion of hESCs and hiPSCs. The advantages of using one condition over the other vary based on the particular experiment for which the hESCs will be utilized. Chemically defined feeder-free conditions may be preferred for chemical screens or other uses in which the media composition must be known. In addition, such conditions eliminate contamination of hESCs or their derivatives with MEFs or MEF-derived molecules such as nonhuman sialic acid (Martin et al., 2005). However, the KOSR/MEF-conditions provide an inexpensive way to expand large numbers of cells suitable for subsequent in vitro differentiation. The authors have successfully transferred hESCs and iPSCs from one condition to the other and vice versa, though some lines grow better in one particular condition.

Critical Parameters bFGF Basic fibroblast growth factor (bFGF, also known as FGF2) is critical to the hESC culture system, as FGF signaling has a central role in sustaining hESCs (Wang et al., 2005). Despite the endogenous expression of bFGF by hESCs, exogenous bFGF is required for efficient subcloning, growth, and inhibition

Embryonic and Extraembryonic Stem Cells

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Supplement 14

A

B

Figure 1C.10.5 Morphologies of hESCs cultured on varying MEF densities. (A) iMEFs plated too sparsely are unable to support proper hESC pluripotency, and cause the periphery of the colony to differentiate. (B) iMEFs plated too densely do not provide hESCs the room required to grow optimally. Scale bar = 250 μm.

of differentiation (Amit et al., 2000; Vallier et al., 2005; Levenstein et al., 2006). Researchers use varying concentrations of bFGF in KOSR/iMEF-based culture systems, ranging from 4 to 20 ng/ml (Lerou et al., 2008). The authors recommend using 10 ng/ml of bFGF, which is optimal for this protocol. As with most growth factors, repeated freeze/thaw cycles should be avoided. MEF quality Quality control testing of the iMEFs is one of the most critical factors for the successful culture of hESCs, since each batch of iMEFs varies in its capacity to support these cells (Amit et al., 2003). When low-quality iMEFs are used, the culture shows increased spontaneous differentiation and poor colony morphology. The optimum plating density of iMEFs is important for minimizing hESC differentiation and sustaining appropriate proliferation. hESC plated on high-density iMEFs will remain small, grow in a more threedimensional dome shape, and start to differentiate at the top of the colonies. At a low density of iMEFs, hESC peripheral differentiation will increase (Fig. 1C.10.5A,B).

Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

Appropriate clump size Regardless of the passaging technique, mechanical or enzymatic, it is important that the hESCs remain in small clumps (100 to 200 cells/clump, though optimal clump sizes vary from line to line), to preserve the replating efficiency and pluripotency of the hESC culture. Plating of clumps that are too small, or of single cells, will drastically diminish plating efficiencies while increasing differentiation. Furthermore, the stresses associated with

complete enzymatic dissociation and single cell plating creates a strong selective advantage for survival and outgrowth of abnormal cells. In contrast, clumps that are too large may not attach properly (e.g., part of the large colony fragment may fold onto itself and form a differentiating appendix in the center of the colony; refer to Figure 1C.1.10B). Large clumps are also more likely to “ball up” and form embryoid-body-like structures that will not attach. The degree to which this problem occurs varies from line to line and also depends on the amount of time the clumps are left in suspension prior to replating (the time from colony detachment and trituration to replating should be kept to a minimum). Appropriate passage timing and plating densities To achieve optimum attachment and continued undifferentiated proliferation, appropriate passage timing is also important to ensure success with this culture system. Passaging cells too early will slow overall expansion rates and may lead to low plating efficiencies. In contrast, letting cultures get too dense and colonies get too large will increase differentiation and metabolic stresses. hESC cultures are typically passaged every 5 to 7 days, depending on the state of the culture and the growth rate of the particular cell line. Optimal passaging time is often judged by percent confluency (usually ∼70% confluent, to ensure the cells are still growing near-exponentially); however, one should never allow a culture to become more than 10% differentiated in the interest of increasing confluency. Large hESC colonies that have not yet begun to differentiate are the best indication that a culture is

1C.10.12 Supplement 14

Current Protocols in Stem Cell Biology

Table 1C.10.1 Troubleshooting Guide for Feeder-Based Culture of hESCs or iPSCs

Problem

Possible cause

Solution

Inefficient detachment during Collagenase IV enzyme solution collagenase IV splitting has low activity

Make sure collagenase IV solution is fresh and has not been stored at elevated temperatures for extended periods of time. Be sure to wash wells with DMEM/F12 prior to adding prewarmed collagenase IV solution.

Low or no attachment after plating

Collagenase IV splitting: enzyme may not have been adequately rinsed/washed away

Wash completely with DMEM/F-12 after collagenase treatment

Inappropriate clump size: cell clumps were smaller than 50 cells/clump

Increase clump size (use larger pipet and reduce triturating forces) or add ROCK inhibitor Y27632 (5 μM) for the first 24 hr after plating

Inappropriate passage timing

Cells should be passaged at peak of growth (mid log-phase)

Passaging procedure too slow Minimize the time from detachment and triturating (clumps kept in suspension for too to plating long) Uneven plating

Slow proliferation or excessive differentiation

Good attachment but proliferation is too fast, no differentiation

Plate (medium) was swirled after plating

Avoid circular movements that may cause the medium to swirl within the well

Cell clumps settled within pipet during plating of multiple wells

Plate one well at a time and mix in between wells

Uneven clump sizes

Be sure to evenly scrape the entire well and to triturate the entire suspension

Poor quality KOSR

Batch test KOSR to ensure sufficient support, then purchase a large order to maintain consistency for 6 months or more

Poor quality MEFs

Test MEF viability and mycoplasma contamination. If necessary, switch to a different batch or source.

Suboptimal MEF density

Optimal MEF density is cell line dependent and may require fine-tuning

Suboptimal clump size

Increase or decrease clump size

Suboptimal passage timing

Even if cells are able to attach after having been passaged too late or too soon, continuously subjecting cells to such treatment will result in increased spontaneous differentiation. See “Inappropriate passage timing” under “Low or no attachment.”

Mycoplamsa contamination

Check mycoplamsa contamination once a month

Abnormal karyotype

Check karyotype of the main stock of each stem cell line and every 10-20 passages thereafter

Other (epi-)genetic abnormalities

These are more difficult to exclude than gross chromosomal abnormalities. Best to re-initiate culture from a low-passage vial. continued Embryonic and Extraembryonic Stem Cells

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Supplement 14

Table 1C.10.1 Troubleshooting Guide for Feeder-Based Culture of hESCs or iPSCs, continued

Problem

Possible cause

Solution

Poor plating efficiency or colony recovery after thaw

Cell clumps too small

Increase clump size for subsequent freezes (use larger pipet and reduce triturating forces) and add ROCK inhibitor Y27632 (10 μM) for the first 24 hr after thawing

Cell clumps too large

Gently triturate clumps during plating

Excessive stress during freezing or thawing

Strictly follow and swiftly execute the protocol

ready to be passaged. Avoid letting colonies touch and fuse with one another, or get too large, as this would slow their growth and may increase differentiation. The best results will be obtained when all clumps are of the same, near-optimal size and are plated evenly throughout the well. Care must be taken to avoid hydrodynamic effects from causing cell clumps to accumulate disproportionally in one area of the well (e.g., causing the cell clump suspension to move around a 6-well in a circular motion by horizontally swirling the plate will concentrate the cell clumps in the center of the well). hESCs cultures are noticeably more “healthy,” in terms of expansion rates and lack of differentiation, when colonies are evenly spaced and neither too dense nor too sparse. When plating multiple wells it must also be taken into account that cell clumps settle much more quickly than single cell suspensions— i.e., aspirating the entire suspension of cell clumps followed by slowly dispensing equal volumes into the new wells will most likely lead to uneven plating densities between the wells.

in teratomas or during differentiation in vitro, although the efficiency of directed differentiation will depend in large part on the robustness of the differentiation protocol and the potential of the particular line. Cells cultured according to the mechanical or bulk methods should proliferate readily, maintain a normal karyotype, and be passaged at split ratios of ∼1:3-1:8 every week. The spontaneous differentiation of a given culture should not exceed 10% but will often vary by hESC line. Assuming cultures are wellmaintained, one confluent well of hESCs can be expanded to more than 100 times its original quantity over the course of four passages (assuming a 1:5 split ratio).

Troubleshooting

MEF preparation Thawing and plating iMEFs will require ∼45 min of preparation. Allow at least 20 min for gelatin-coated plates to incubate at 37◦ C. The appropriate amount of iMEF medium should also be prewarmed for 15 to 30 min prior to use. Thawing and plating the iMEFs will take ∼15 min, but could take longer if many plates are being handled simultaneously.

See Table 1C.10.1 for troubleshooting suggestions.

Anticipated Results

Clump Passaging and Expansion of hES and iPS Cells on MEF Feeder Cells

This protocol is designed to generate large numbers of hESCs for experimental manipulation. If hESCs were thawed and passaged according to these protocols, they will maintain classic embryonic stem cell morphology including high nucleus-to-cytoplasm ratio, tight colony borders, and a homogeneous cell appearance within hESC colonies. Pluripotency markers including TRA-1-60, Oct4, and Nanog will be highly expressed at any point throughout the culture, but will also become rapidly down regulated upon differentiation. The cells will maintain their ability to differentiate into all three germ layers

Time Considerations Reagent and media preparations Allow 15 to 20 min for the preparation of sterile stock solutions including bFGF, collagenase IV, and 0.1% BSA in CMF-DPBS. Media recipes require only 10 min to prepare and filter.

Passaging iMEF-coated plates should be prepared the day before usage, or a few (2 to 3) days in advance; fresh iMEFs are highly recommended for thawing hESCs. Prewarming aliquots of DMEM/F-12 and hESC medium is recommended at least 15 to 30 min prior to passaging. Cells should be supplemented with onehalf volume of prewarmed hESC medium at

1C.10.14 Supplement 14

Current Protocols in Stem Cell Biology

least 1 hr prior to passaging. The prepassage feed ensures the hESCs are in a growth phase when passaged. The entire process of passaging, from removing differentiation to replating, can be achieved in as little as 20 min per plate. However, the rate-limiting step is the amount of time it takes to remove gross differentiation from your hESC culture, which should be done slowly and carefully. Routine culture Medium should be warmed to 37◦ C ∼15 to 30 min before routine medium changes. Once medium is warmed, medium changes should take 3 min or less per plate.

Literature Cited Amit, M., Carpenter, M.K., Inokuma, M.S., Chiu, C.P., Harris, C.P., Waknitz, M.A., ItskovitzEldor, J., and Thomson, J.A. 2000. Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227:271278. Amit, M., Margulets, V., Segev, H., Shariki, K., Laevsky, I., Coleman, R., and Itskovitz-Eldor, J. 2003. Human feeder layers for human embryonic stem cells. Biol. Reproduct. 68:2150-2156. Draper, J.S., Smith, K., Gokhale, P., Moore, H.D., Maltby, E., Johnson, J., Meisner, L., Zwaka, T.P., Thomson, J.A., and Andrews, P.W. 2004. Recurrent gain of chromosomes 17q and 12 in cultured human embryonic stem cells. Nat. Biotechnol. 22:53-54. Lerou, P.H., Yabuuchi, A., Huo, H., Miller, J.D., Boyer, L.F., Schlaeger, T.M., and Daley, G.Q. 2008. Derivation and maintenance of human embryonic stem cells from poor-quality in vitro fertilization embryos. Nat. Protoc. 3:923-933.

Levenstein, M.E., Ludwig, T.E., Xu, R.H., Llanas, R.A., VanDenHeuvel-Kramer, K., Manning, D., and Thomson, J.A. 2006. Basic fibroblast growth factor support of human embryonic stem cell self-renewal. Stem Cells 24:568-574. Liu, Y., Song, Z., Zhao, Y., Qin, H., Cai, J., Zhang, H., Yu, T., Jiang, S., Wang, G., Ding, M., and Deng, H. 2006. A novel chemicaldefined medium with bFGF and N2B27 supplements supports undifferentiated growth in human embryonic stem cells. Biochem. Biophys. Res. Commun. 346:131-139. Ludwig, T.E. and Thomson, J.A. 2009. Defined, feeder-independent medium for human embryonic stem cell culture. Curr. Protoc. Stem Cell Biol. 2:1C.2.1-1C.2.16. Ludwig, T.E., Bergendahl, V., Levenstein, M.E., Yu, J., Probasco, M.D., and Thomson, J.A. 2006. Feeder-independent culture of human embryonic stem cells. Nat. Methods 3:637646. Martin, M.J., Muorti, A., Gage, F., and Varki, A. 2005. Human embryonic stem cells express an immunogenic nonhuman sialic acid. Nat. Medicine 11:228-232. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Vallier, L., Alexander, M., and Pedersen, R.A. 2005. Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell Sci. 118:4495-4509. Wang, L., Li, L., Menendez, P., Cerdan, C., and Bhatia, M. 2005. Human embryonic stem cells maintained in the absence of mouse embryonic fibroblasts or conditioned media are capable of hematopoietic development. Blood 105:45984603.

Embryonic and Extraembryonic Stem Cells

1C.10.15 Current Protocols in Stem Cell Biology

Supplement 14

Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

UNIT 1C.11

Allen K. Chen,1 Xiaoli Chen,1 Andre B.H. Choo,1 Shaul Reuveny,2 and Steve K.W. Oh1 1

Stem Cell Group, Bioprocessing Technology Institute, A*STAR (Agency for Science, Technology and Research), Centros, Singapore 2 Department of Biotechnology, Israel Institute for Biological Research, Ness Ziona, Israel

ABSTRACT This unit describes the routine maintenance and expansion of undifferentiated human embryonic stem cells (hESC) on cellulose microcarriers. Conventionally, hESCs have been maintained on feeder cells or extracellular matrix–coated two-dimensional tissue culture plates. The expansion of hESC on a tissue culture platform is limited by the available surface area and the requirement of repetitive subculturing to reach the required cell yield. Here, we show that expansion of hESC can be carried out in a three-dimensional suspension culture using Matrigel-coated cellulose microcarriers. hESCs from a tissue culture plate can be seeded directly onto the microcarriers; hESC microcarrier culture is passaged and expanded by mechanical dissociation of the cells without enzyme. Expansion of the culture in a 100-ml spinner ßask is also described. Long-term culture of hESC on the microcarriers maintains typical pluripotent markers (OCT-4, Tra-1-60, and SSEA-4) and stable karyotype. Spontaneous differentiations of microcarrier-maintained hESCs in vitro (embryoid body formation) and in vivo (teratoma formation in SCID mouse) have demonstrated formation of the three germ layers. These protocols can also be applied equally well to human induced pluripotent stem cells. Curr. Protoc. Stem Cell C 2010 by John Wiley & Sons, Inc. Biol. 14:1C.11.1-1C.11.14.  Keywords: human embryonic r stem cells r hESCs r microcarriers r hESC expansion r induced pluripotent stem cells r iPSC

INTRODUCTION This unit describes the routine maintenance and expansion of undifferentiated human embryonic stem cells (hESC) on cellulose microcarriers. The development of a microcarrier platform for the cultivation of undifferentiated hESC allowed the expansion of hESC in three-dimensional suspension cultures using conventional stirred cultures. Studies have demonstrated that microcarrier-based hESC cultivation achieved higher cell density than the conventional tissue culture plate while maintaining all the characteristics of undifferentiated hESC (Lock and Tzanakakis, 2009; Nie et al., 2009; Oh et al., 2009). The protocols presented below are detailed descriptions of those published by Oh et al. (2009). The cellulose carriers are cylindrical in shape (length 130 ± 60 μm × diameter 35 ± 7 μm) and positively charged. This unit begins with the preparation of Becton Dickinson Matrigel-coated cellulose microcarriers, followed by the seeding of hESC onto the microcarriers, passage of microcarrier culture, and culture expansion using spinner ßask. The protocol can ultimately be used as the basis for larger-scale bioreactors. NOTE: The following procedures are to be performed in a Class II biological safety cabinet. NOTE: Aseptic technique is required for handling of all the solutions and equipments in contact with living cells. Current Protocols in Stem Cell Biology 1C.11.1-1C.11.14 Published online September 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01c11s14 C 2010 John Wiley & Sons, Inc. Copyright 

Embryonic and Extraembryonic Stem Cells

1C.11.1 Supplement 14

BASIC PROTOCOL 1

SEEDING hESCs FROM TWO-DIMENSIONAL CULTURE ONTO MATRIGEL-COATED CELLULOSE MICROCARRIERS The protocol for the initiation of hESC microcarrier culture from two-dimensional cultures is described below. The hESC culture used for seeding should be of high quality with at least 80% of the cells expressing the pluripotent marker TRA-1-60.

Materials hESC (HES-3 and HES-2; ES International), expanded colonies cultured on Matrigel-coated dish Conditioned hESC growth medium (see recipe) Matrigel-coated cellulose microcarriers (Support Protocol 2) Stereomicroscope 1-ml and 200-μl pipettor and sterile-Þltered tips StemPro EZPassage Disposable Stem Cell Passaging Tool (Invitrogen, cat. no. 23181-010) Cell scraper, 24 cm (TPP, cat. no. 99002) NucleoCounter (ChemoMetec A/S, NucleoCounter SCC-100) Orbital shaker (e.g., IKA Mixing Oribital Shaker KS260 Control, cat. no. 2980300) 37◦ C humidiÞed incubator with 5% CO2 1. Examine the conßuent hESC culture on Matrigel-coated 60 × 15–mm tissue culture dish under a stereomicroscope. Remove differentiated regions within each colony using sterile-Þltered 200-μl tips attached to a pipettor. Coat the dish with 4 ml of Matrigel stock solution (BD Biosciences) diluted 1:30 in cold unconditioned hESC growth medium overnight at 4◦ C (BD Biosciences, cat. no. 353004).

2. Replace the medium of the culture with 4 ml fresh conditioned hESC growth medium. 3. Using a StemPro EZPassage Disposable Stem Cell Passaging Tool, cut the colonies on the culture dish according to the manufacturer’s instruction. This step should be carried out under a stereomicroscope inside a Class II biological safety cabinet. The StemPro EZPassage Disposable Stem Cell Passaging Tool is the preferred tool in this protocol for manual passaging of hESC because of its simplicity for cutting colonies in a short time. In brief, the tool consists of a handle with a roller attached. As the tool is rolled across the conßuent culture dish, the grooves on the roller cut hESC colonies with equal spacing. Then the dish is rotated 90◦ and the colonies are cut again with the tool. The resulting uniform-cut-size colonies are generated as shown in Figure 1C.11.1C.

4. Dislodge the sliced colonies from the plate by using a cell scrapper (Fig 1C.11.1D). We have found cell clumps generated from the passaging tool were uniform in size. Seeding the cells obtained by enzymatic dissociation is also possible (UNIT 1C.1).

5. Take two samples of 200 μl each to determine the viable cell concentration of cell clumps using the NucleoCounter. Adjust the cell concentration to 8 × 105 cells/ml by adding more conditioned hESC growth medium.

Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

NucleoCounter is a cell-counting instrument based on the staining of ßuorescent dye propidium iodide (PI) on DNA of nuclei. In brief, the cell sample is added with an equal volume of the lysis buffer Reagent A, which disrupts the plasma membrane of the cells and exposes the nuclei for staining. Furthermore, Reagent A works effectively in disaggregating the cell clumps and the cells on microcarriers. The stabilizing Reagent B is then added into the sample in an equal volume as Reagent A to allow for PI staining. The Þnal cell sample mixture is loaded into a cassette at ∼50 μl. The cassette (NucleoCassette) consists of a ßow channel, which allows only nuclei to pass into a chamber of immobilized PI to stain them, and then they move on to a measurement chamber. The instrument reads the

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numbers of nuclei and provides the cell count. Dead cells from the supernatant can be measured by loading the sample directly into the cassette without any treatment to determine percent cell viability. Microcarrier fragments do not interfere with nuclear staining by PI. Instead of using the NucleoCounter, other standard nuclear count methods (Sandford et al., 1951; White and Ades, 1990; Lin et al., 1991; Shah et al., 2006; Altman et al., 2008) can be used or hESC clumps can be dissociated into single cells using dissociating enzymes and viable cells quantiÞed using the trypan-blue-exclusion method (UNIT 1C.3).

6. Seed 1 ml of cut cell clumps at 8 × 105 cells/ml into the wells of a 6-well plate containing 4 ml of Matrigel-coated cellulose microcarriers at a concentration of 4 mg/ml. This is equivalent to a starting cell density of 1.6 × 105 cells/ml. Non-tissue culture polystyrene 6-well plates are not recommended as we have encountered cell attachment on to the plate.

7. Place the plate onto an orbital shaker in a 37◦ C humidiÞed incubator with 5% CO2 and agitate at 110 rpm for 2 hr to allow cell attachment to microcarriers (immediate cell microcarrier aggregation can be seen). In the authors’ experience, the clustering of the cells and microcarriers in the center of the well enhanced cell attachment to the microcarriers.

8. Perform daily medium changes by gently tilting the plate to one side, allowing cellmicrocarrier aggregates to settle and remove 4 ml of spent medium. Add 4 ml of fresh conditioned hESC growth medium into the well. 9. Passage the microcarrier culture once a week to ensure that cells are not too conßuent on the microcarriers and to avoid adverse medium condition, in particular the decrease in pH and accumulation of waste metabolites like lactic acid and ammonium.

PREPARATION OF CELLULOSE MICROCARRIERS This protocol describes the preparation of cellulose microcarriers for the cultivation of hESC in suspension. The authors have tried three different cellulose microcarriers (DE52, DE53, and QA52) from Whatman. Even though these cellulose microcarriers differ in their surface charges, the authors found no signiÞcant differences in term of cell growth and pluripotent marker expression.

SUPPORT PROTOCOL 1

Materials Cellulose carriers (DE52, DE53, or QA52; Whatman, cat no. WH/CH/CO/4057-050, WH/CH/CO/4058-050 or WH/CS/OO/4065-050) 4 M HCl Dulbecco’s phosphate-buffered saline without calcium chloride and magnesium chloride (DPBS− ; Invitrogen, cat. no. 14190-144) Weighing balance 500-ml glass bottles coated with Sigmacote (Sigma-Aldrich, cat. no. SL2) 1-ml pipettor and tips pH meter 50-ml serological pipet Steam sterilizer 1. Add 25 g of preswollen cellulose microcarriers into the precoated 500-ml glass bottle containing 500 ml DPBS− . The protocol describing coating of glassware with Sigmacote is given in the manufacturer’s product information sheet. In brief, 2 ml of Sigmacote is applied directly onto the dry inner surface of the 500-ml glass bottle. Make sure that all surfaces are covered by pipetting with 5-ml pipet and rotating the bottle. After removing excess Sigmacote, air dry the bottle overnight and rinse with water. The bottle is then sterilized by autoclaving.

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2. Maintain the microcarriers in suspension by shaking the bottle in circular motion. At the same time, measure the pH of the solution. The pH should be alkaline, above pH 8. A magnetic stirrer is not recommended, as it can grind and break up the microcarriers.

3. Add 4 M HCl dropwise to a Þnal stabilized pH level of 7.2. 4. Allow the solution to stand for at least 30 min, during which the microcarriers settle at the bottom of the bottle. 5. Remove 80% to 90% of solution without microcarriers and replace it with an equal volume of DPBS− using 50-ml serological pipettes. 6. Steam sterilize the microcarrier suspension (121◦ C for 20 min). SUPPORT PROTOCOL 2

COATING MICROCARRIERS WITH MATRIGEL Becton Dickinson’s (BD) Matrigel is considered to be critical for culturing of hESC on microcarriers. Our group, as well as others, showed that seeding hESC on uncoated microcarriers usually resulted in limited cell growth or undesired spontaneous differentiation. Instead of BD Matrigel, other extracellular matrix (ECM) components, such as laminin, can also be considered to replace Matrigel in order to have deÞned coatings on the cellulose microcarriers. This protocol describes the preparation of a whole 6-well plate for the cultivation of hESC on microcarriers. One plate would require 120 mg of Matrigel-coated cellulose microcarriers (20 mg per well). NOTE: Matrigel stock solution should be divided into 1-ml aliquots per 1.5-ml sterile tube and stored up to 1 month at −20◦ C. An aliquot of Matrigel is thawed overnight at 4◦ C prior to use. In order to prevent solidiÞcation of Matrigel, tips and tubes used in the preparation should be cooled in the benchtop cooler (StrataCooler LP Benchtop Cooler; Agilent Technologies, cat. no. 401349) or on ice before use.

Materials Sterile cellulose microcarrier suspension (50 mg/ml) in Dulbecco’s phosphate-buffered saline solution (see Support Protocol 1) Unconditioned hESC medium (see recipe) BD Matrigel Basement Membrane Matrix (Beckon Dickson, cat. no. 354234) Conditioned hESC growth medium (see recipe), cold 1.5-ml and 50-ml centrifuge tubes, sterile 5-ml and 25-ml serological pipets, sterile 1-ml sterile Þltered tips, pre-cooled Laboratory platform rockers Ultra-low adherent 6-well plate (Corning, cat. no. 3471) 37◦ C humidiÞed incubator with 5% CO2 1. Dispense 2.4 ml of the sterile microcarrier suspension in PBS prepared previously into a 50-ml tube using a 5-ml sterile serological pipet and keep on ice. 2. Dispense 21.6 ml of cold unconditioned hESC medium (dilution medium) into the tube containing microcarriers using a 25-ml sterile serological pipet. Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

3. Add 0.8 ml of thawed Matrigel using a cold, sterile 1-ml Þltered tip to the 50-ml tube containing the cold dilution medium and microcarriers. Mix well by gently inverting the 50-ml tube horizontally onto a rocker for gentle mixing for at least 4 hr before use.

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If not for immediate use, the coated microcarriers can be placed onto the rocker overnight at 4◦ C. It is also possible to store the Matrigel/microcarrier mixture at 4◦ C for at least one month for future use.

4. Allow the microcarriers to settle at the bottom of the tube. Aspirate the solution above the microcarrier. Centrifugation at a low speed of 200 × g for 1 min can be used to shorten the microcarrier settling time.

5. Wash the microcarriers with 24 ml of unconditioned hESC medium. Allow the microcarrier to settle and remove the solution above the microcarriers.

Figure 1C.11.1 Phase-contrast microscopy of microcarriers and hESC at various stages of the protocols. Scale bar = 200 μm. (A) Uncoated cellulose microcarriers in hESC growth medium. (B) BD Matrigel-coated cellulose microcarriers in hESC growth medium. (C) hESC colonies on 60 × 15–mm Matrigel-coated dish; the culture has been sliced into a grid motif using a StemPro EZPassage Disposable Stem Cell Passaging Tool (Invitrogen). (D) The sliced pieces of hESC colonies have been lifted up into suspension before seeding to microcarriers. (E) Day 6 images of hESC microcarrier static culture showing the formation of large cell/microcarrier aggregates. (F) Day 7 images of hESC microcarrier culture in a spinner ßask with smaller and denser cell/microcarrier aggregates.

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6. Add 24 ml conditioned hESC growth medium and aliquot 4 ml into each well of the ultra-low adherent 6-well plate. Place the plate in a 37◦ C humidiÞed incubator with 5% CO2 for at least 1 hr before use. Matrigel-coated microcarriers have a tendency to form aggregates by themselves, whereas this is not seen in uncoated microcarrier suspensions. Hence by making this observation, one can determine the quality of Matrigel coating (see Fig. 1C.11.1A,B to compare the differences between Matrigel-coated and uncoated microcarriers). BASIC PROTOCOL 2

PASSAGING hESCs FROM MICROCARRIERS TO MICROCARRIERS This protocol has been routinely used by our group to maintain hESC on microcarriers for at least 25 weeks (Oh et al., 2009). Suspension microcarrier hESC cultures have been found to have a stable karyotype, a consistent percentage of cells expressing pluripotent markers, and the cells are able to undergo spontaneous differentiation to three germ layers both in vitro and in vivo. hESC cultures that have been transferred back to a Matrigelcoated dish appear indistinguishable from those maintained routinely on Matrigel-coated dishes, in terms of cell morphology (see Fig. 1C.11.2). Using the hESC microcarrier passaging method, it is simple to expand hESC or to generate inoculums for scale-up in a spinner ßask or bioreactor (Basic Protocol 3).

Materials hESC on Matrigel-coated microcarriers in one well of an ultra-low adherent 6-well plate (see Support Protocol 2) Conditioned hESC growth medium (see recipe) 1-ml and 200-μl pipettors with sterile-Þltered tips Sterile 5-ml serological pipet NucleoCounter (ChemoMetec A/S, NucleoCounter SCC-100) Ultra-low adherent 6-well plate (Corning, cat. no. 3471) BD Matrigel Basement Membrane Matrix (Beckon Dickson, cat. no. 354234) Sterile cellulose microcarrier suspension (50 mg/ml) in phosphate-buffered saline solution (Support Protocol 1) Orbital shaker (e.g., IKA Mixing Oribital Shaker KS260 Control, cat. no. 2980300) 37◦ C humidiÞed incubator with 5% CO2

Figure 1C.11.2 Plating of an hESC-microcarrier aggregate (arrow) onto a Matrigel-coated dish, after 1 day (A) and 3 days (B). Scale bar = 200 μm.

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Perform continuous cultivation of hESC on microcarrier 1. Prepare hESC on Matrigel-coated microcarriers according to Support Protocol 2. 2. Exchange the medium of the microcarrier culture to be passaged with 4 ml conditioned hESC growth medium. 3. Break up aggregates of the hESC microcarrier culture using a 1-ml Þltered pipet tip by continuously aspirating and dispensing until an even distribution of cell/microcarrier clumps is achieved (200 to 300 μm in size). Figure 1C.11.1E shows the image of large cell/microcarrier aggregates after 6 days of cultivation. This step should be carried out under a stereomicroscope inside a Class II biological safety cabinet. Enzymatic dissociation of cells from microcarriers for further passaging has been used by our group. We found that the use of enzyme could cause a decrease in viable cell concentration. No signiÞcant differences in cell yields of cultures seeded with inoculums from enzymatic or mechanically treated cells were observed. However, the mechanically treated cell culture usually had higher number of cells expressing pluripotent markers compared to the enzymatically treated one.

4. Measure the total volume of medium in each well of the 6-well plate using a sterile 5-ml serological pipet. We have observed the loss of medium due to evaporation and pipet volume error from medium exchange. For accurate reporting on cell yield at the end of a passage, it is important to take note of the actual culture volume per well to calculate the total number of cells.

5. Remove 200 μl from each well for cell count using a NucleoCounter. An extra 200-μl sample can be used to determine the nonviable cell concentration using the NucleoCounter. However, we found it is not necessary, as most of the cells on microcarriers are viable (>90%). Nonviable cells in suspension are mostly removed during the medium exchange step. Another method for measuring cell concentration on microcarriers is by nuclear staining with crystal violet solution (0.2% w/v crystal violet in 0.2 M citric acid solution) according to a study by Hu and Wang (1987). In brief, the 200-μl hESC microcarrier sample was added to a 200-μl crystal violet solution. After incubation for at least 30 min at 37◦ C, use a 1-ml pipet to carry out repetitive aspirating and dispensing to release the nuclei from the microcarriers. The stained nuclei are then counted using a hemacytometer.

6. Seed the cell-microcarrier aggregates into a new well with newly coated microcarriers. Top up medium volume with conditioned hESC growth medium to 5 ml per well. Seeding density for HES-3 and HES-2 in our study is 4 × 105 cells/well, equivalent to 0.8 × 105 cells/ml. During the Þrst few hours after seeding, many small aggregates of microcarriers coated with cells are generated. Thereafter, during cell propagation, hESC are able to grow onto adjacent cell-free microcarriers, as well as increase in number on existing microcarriers, under static conditions. The authors do not typically passage by split ratios but prefer to seed cells at the same cell density at each passage. Typically, expansion is about 10-fold every week (expansion less than 5-fold or doubling time longer than 35 hr is considered as poor cell growth). This might be due to poor Matrigel coating or onset of spontaneous differentiation.

7. Place the newly seeded plate onto an orbital shaker in a 37◦ C humidiÞed incubator with 5% CO2 and agitate at 110 rpm for 2 hr.

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The remaining unseeded cells can be used for ßow cytometry, RNA for qRT-PCR, and protein for immunoblots. Dissociating enzymes can be used to detach cells from microcarriers for applications like ßow cytometry, which require single-cell suspensions without microcarriers. Hence, in order to remove microcarriers from suspension cells, we used a 40-μm cell strainer (BD Biosciences, cat. no. 352340) to Þlter out the microcarriers. Similarly, this procedure should also be applied when extracting RNA for gene expression study as RNA can bind to the microcarriers and might hinder the subsequent puriÞcation steps. Removal of cells from microcarriers is not necessary for extraction of proteins from the cells.

8. Feed the cells every day by removing 80% of the spent medium (∼4 ml) and adding equal volume of fresh conditioned hESC growth medium. 9. Continuously passage the hESC microcarrier culture for at least 5 passages. For karyotype analysis, hESC microcarrier culture is plated on Matrigel coated 60×15– mm tissue culture dish, where the removal of microcarriers is preferred. Similarly, plated hESC is also used for teratoma formation in mouse with severe combined immunodeÞciency (SCID). The authors usually observe the plated cells would spread and expand around the cell-microcarrier aggregates (Fig. 1C.11.2). Their morphologies are indistinguishable from those cultured routinely on Matrigel-coated tissue culture dish. BASIC PROTOCOL 3

EXPANSION OF hESC USING A SPINNER FLASK The following protocol describes the expansion to a 50-ml hESC microcarrier culture in a 100-ml spinner ßask. Even though a larger vessel has not yet been tested, we believe the current method can be easily adapted by adjusting the stirrer position and/or speed to ensure suspended microcarriers in the medium while avoiding shear stress to hESCs.

Materials hESC (HES-3 and HES-2, ES International), conßuent and adapted (more than 4 passages) to microcarrier culture (Basic Protocol 1) Conditioned hESC growth medium (see recipe) Ultra-low-adherent 6-well plate (Corning, cat. no. 3471) 100-ml spinner ßasks (Bellco, cat. no. 1965-00100) coated with Sigmacote (Sigma-Aldrich, cat. no. SL2) Autoclave 37◦ C humidiÞed incubator with 5% CO2 Orbital shaker (e.g., IKA Mixing Oribital Shaker KS260 Control, cat. no. 2980300) 1-ml and 200-μl pipettors with sterile-Þltered tips NucleoCounter (ChemoMetec A/S, NucleoCounter SCC-100) Stereomicroscope Magnetic stirrer (e.g., Thermolyne Cellgro Stirrer, Thermal ScientiÞc Barnstead, cat. no. S45600) Sterile 5-ml serological pipets Sterile 15- and 50-ml centrifuge tubes Additional reagents and equipment for preparing microcarriers coated with Matrigel (Support Protocol 2) Prepare the cells 1. At a time point 7 days before seeding the spinner ßask, inoculate three wells of a 6-well plate at viable cell density of 0.8 × 105 cells/ml (5 ml medium per well). Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

Three wells should be sufÞcient to generate 1×107 viable cells.

2. At a time point 1 day before seeding the spinner ßask, prepare 200 mg microcarriers coated with Matrigel in a sterile 50-ml centrifuge tube (Support Protocol 2).

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3. Sterilize a 100-ml pre-coated spinner ßask using an autoclave. 4. At a time point 1 hr before seeding the spinner ßask, replace the microcarrier-Matrigel solution in step 2 with 15 ml conditioned hESC growth medium. 5. Transfer 15 ml microcarrier suspension in the conditioned hESC medium to the spinner ßask and then place the spinner ßask onto the magnetic stirrer plate at a speed of 25 rpm in a 37◦ C humidiÞed incubator with 5% CO2 . 6. For the preparation of the seeding culture, using a 1-ml pipet, mechanically break up the cell/microcarrier aggregates into ∼200 to 300 μm in size and determine the viable cell concentration with a NucleoCounter as described in Basic Protocol 1 (step 5). This step should be carried out under a stereomicroscope inside of a Class II biological safety cabinet.

Transfer the cells/microcarriers to spinner ßask 7. Transfer 1 × 107 viable cells on microcarriers, cultured previously in step 1, into the spinner ßask and top up medium volume to 25 ml with conditioned hESC growth medium. 8. Leave the spinner ßask in a static condition inside the incubator for 24 hr. 9. After 1 day, increase the medium volume to 50 ml with conditioned hESC growth medium 10. Place the spinner ßask on a stirring platform inside the incubator and set the speed to 25 rpm. The microcarriers by themselves are not affected by high agitation speed (up to 100 rpm). However, it is important to use the lowest speed possible to suspend microcarriers, as higher speeds can cause cell detachment, low viability, and eventually cessation in cell growth.

Count the cells 11. For cell concentration determination, remove the spinner ßask from the incubator, place it inside a biological safety cabinet, and take a 2-ml sample using either a 5-ml or 10-ml serological pipet. 12. Determine the cell count using a NucleoCounter. It is important to keep the microcarrier culture in suspension homogeneously before sampling. Smaller sampling volumes might affect the accuracy of the measured cell concentration. Due to the agitation, the cell/microcarrier aggregates are usually more uniform and smaller than those observed in static microcarrier culture (Fig. 1C.11.1F).

Feed the cells 13. Prewarm fresh conditioned hESC growth medium in a 37◦ C incubator or water bath. 14. Leave the spinner ßask in a static condition. Once cell-microcarrier aggregates have settled down to the bottom of the spinner ßask, remove the culture medium with a 10-ml serological pipet without causing much disturbance to the cell-microcarrier aggregates. Larger-volume pipets are too large for the spinner ßask openings. The collected spent medium in 50-ml tubes can be used for further analyses such as pH, osmolarity, and metabolite concentrations.

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15. Add in an equal volume of fresh conditioned hESC growth medium to the spinner ßask. 16. Optional. Take a 2-ml sample using either a 5-ml or a 10-ml serological pipet for cell count and other analyses. For metabolism studies, it is important to take another sample after medium exchange. Hence, the daily consumption rates of metabolites can be calculated.

17. Replace 80% of the spent medium with fresh hESC conditioned growth medium daily throughout the entire propagation period. Spinner ßasks seeded at 2 × 105 cells/ml grow for 5 days reaching density of at least 2 × 106 cells/ml.

18. When the culture reaches plateau, harvest cells for further expansion or differentiation. For further expansion, cells from the spinner ßask can be used to seed another spinner ßask or bioreactor with larger volume to which fresh new Matrigel-coated microcarriers are added using the same seeding density of 2 × 105 cells/ml and microcarrier concentration of 4 mg/ml. For differentiation, embryoid bodies can be generated by exchanging the culture medium with differentiation medium. The need for cell dissociation from the microcarriers depends on the speciÞc differentiation protocols. It is our observation that keeping the cells on microcarriers is beneÞcial for differentiation, as the cells maintain high viability and generate cell/microcarrier aggregates with more controlled size distribution. These parameters can be important for efÞcient cell differentiation, such as cardiomyocytes.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Conditioned hESC growth medium hESC growth medium is conditioned with mitotically inactivated mouse embryonic Þbroblast (UNIT 1C.1) seeded at a density of 2.4 × 105 cells/ml (8 × 105 cells/cm2 ) in MEF medium (see recipe in UNIT 1C.1). After an overnight incubation in a 37◦ C/5% CO2 humidiÞed incubator, the medium is completely replaced with an equal volume of fresh unconditioned hESC growth medium at room temperature for the generation of the conditioned medium. The process is repeated for 7 days. The collected conditioned hESC growth medium is then Þltered (0.2-μm pore size) and spiked with 10 ng/ml basic Þbroblast growth factor. Conditioned hESC growth medium is used for all steps involving hESC.

Unconditioned hESC growth medium

Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

Knockout D-MEM medium (Invitrogen, cat. no. 10829-018) containing: 15% (v/v) Knockout Serum Replacement (Invitrogen, cat. no. 10828-028) 1% (v/v) Non-essential-amino acids 1 mM L-glutamine 1× penicillin/streptomycin 0.1 mM 2-mecaptoethanol 10 ng/ml basic Þbroblast growth factor Store up to 2 weeks at 4◦ C Unconditioned hESC growth medium is only used when preparing the Matrigel coating on the microcarriers.

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COMMENTARY Background Information Human embryonic stem cells (hESC) are a potential source of cells for future cell therapy applications, as they have the ability to differentiate into multilineage cell types from the three germ layers. In order for hESCs to fulÞll their potential, several key bottlenecks have to be resolved. These include expansion of hESC, differentiation to speciÞc cell lineages, isolation of target cell types, and cryopreservation prior to administration. According to a study by Mummery (2005), in order to treat an infarcted heart, it will require 5 × 109 undifferentiated hESC to generate sufÞcient cardiomyocytes, assuming differentiation efÞciency of 100%. Hence, the actual number of hESC required will be higher considering the poor efÞciency of the current differentiation protocols (Thomas et al., 2009). Ever since hESC were Þrst isolated from the inner cell mass of the blastocyst, an earlystage embryo, they have been routinely cultured on mouse embryonic Þbroblasts (MEF; Thomson et al., 1998; Reubinoff et al., 2000). In order to avoid contamination with mouse feeder cells, there were studies demonstrating the use of human feeders substituting mouse ones for expansion of hESC (Richards et al., 2002; Choo et al., 2004). Later, a feederfree platform was developed, replacing feeders with extracellular matrices such as Matrigel, Þbronectin, laminin, and vitronectin (Xu et al., 2001; Richards et al., 2002; Choo et al., 2006; Skottman and Hovatta, 2006; Braam et al., 2008). Several groups have published methods using enzymatic dissociation method to further simplify hESC expansion in the tissue culture ßask and replacing manual cutting. The enzymatic method has shown to be suitable for maintenance of the undifferentiated state, pluripotency, and stable karyotype. (Hasegawa et al., 2006; Ellerstr¨om et al., 2007; Bajpai et al., 2008), even though there are concerns that the enzymatic method might only be best to apply for limited passages (Baker et al., 2007). However, despite several improvements made on the hESC cultivation protocol, the overall procedure is still labor intensive. An automated two-dimensional culture system handling multiple tissue culture ßasks for scale-up of hESC production was suggested by Thomas et al. (2009); however, capital investment for this machine is high and it might not be an economically viable option to other groups.

Since hESCs cannot be cultured as a singlecell suspension and are adherent-dependent (Baker et al., 2007; Watanabe et al., 2007), we have explored culturing hESCs on microcarriers. Traditionally, methods for expansion of adherent cell line have been based on microcarriers. The microcarrier platform provides a large surface area for cell growth in a homogeneous suspension culture system (Malda and Frondoza, 2006). Currently, microcarriers have been used for vaccine and recombinant protein production (GE Healthcare). Recently, there are several groups who have reported the use of microcarriers for culturing hESCs in suspension. They have demonstrated that the microcarrier platform is superior to the two-dimensional tissue culture ßask–based system in terms of cell growth yield, robustness, and scalability. hESC from microcarrier cultures are shown to be able to maintain stable karyotypes, and remain undifferentiated, and pluripotent (Phillips et al., 2008; Oh et al., 2009). Further studies have demonstrated the feasibility of applying directed differentiation of hESCs, both in situ on the microcarriers, as well as after harvesting and plating in twodimensional cultures (Phillips et al., 2008; Lock and Tzanakakis, 2009; Oh et al., 2009).

Critical Parameters and Troubleshooting hESCs cultured on microcarriers in a static or agitated platform are mainly inßuenced by the quality of cells in term of pluripotent marker expression. Other hESC lines that normally require extensive manual removal of differentiated colonies are usually not suitable for this platform due to the difÞculties in the removal of those spontaneously differentiated cells on the microcarriers. Hence, it is recommended to carry out a long-term stability study (for at least 5 passages) to examine the cell growth yields and the expression of pluripotent markers prior to bulk expansion before differentiation studies. It is also recommended to have a parallel two-dimensional culture of hESCs as a control to the microcarrier cultures. Common problems and solutions encountered during the cultivation of hESC microcarrier cultures can be found in Table 1C.11.1. In this protocol, we have described the use of the StemPro EZPassage Disposable Stem Cell Passaging Tool to generate uniform cell clumps for seeding of microcarriers. Other hESC passage protocols used by the authors or other groups using dissociating enzymes

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Table 1C.11.1 Troubleshooting Guide for Common Problems Faced during hESC Cultivation on Microcarriers

Problem

Possible cause

Solution

Poor cell growth in static microcarrier culture

Poor Matrigel coating on microcarriers

Repeat coating with a different lot of Matrigel

Poor initial cell attachment

Check control organ culture dish for cell attachment Repeat coating with a different lot of Matrigel

Low inoculum concentration

Check accuracy of inoculum concentration Increase inoculum concentration

Inoculating with too small cell clumps

Increase the size of the cut cell clumps

Onset of spontaneous differentiation

Restart microcarrier culture if the expression of pluripotent markers is continuously decreasing in subsequent passages

Low microcarrier concentration

Increase the microcarrier concentration at the start of the culture Add more microcarriers to the existing culture

Poor or decrease in cell attachment

Decrease the stirring speed Increase the length of static attachment period Check the density and condition of inoculum

Shear stress

Decrease the stirring speed

Onset of spontaneous differentiation

Restart the spinner ßask culture with new cultures

Over conßuence in the microcarrier culture

Shorten the cell passage time Increase the microcarrier concentration Increase the frequency of the medium exchange

Quality of seeding culture

Prepare a fresh seeding culture with a high expression of pluripotent markers

Poor cell growth in a spinner ßask

Decrease in pluripotent marker expression

Using enzymes to dissociate cells Use the manual breakage of clumps to seed from microcarrier new cultures Medium quality

Expansion of Human Embryonic Stem Cells on Cellulose Microcarriers

can also be adapted and used. It is important to monitor cell attachment to the microcarriers during the Þrst days after seeding. We have seeded single-cell suspensions obtained from Matrigel-coated culture dishes by enzymatic dissociation with Accutase (Invitrogen, cat. no. A11105-01) onto Matrigel-coated microcarriers. At least 80% of the seeded cells attach to the microcarriers, and conßuent microcarrier cultures were achieved within 7 days (see Fig. 1C.11.1E). It is also possible to use other microcarriers instead of the cellulose microcarriers described in this protocol. The amount of coating Matrigel can be estimated based on the amount added and the total surface area of the microcarriers, followed by examination of the cell growth. It is always our observation

Check the medium preparation method, especially the concentration of the basic Þbroblast factor

that hESCs have a preference to attach and grow on Matrigel-coated microcarriers instead of the uncoated ones. Cultivation of hESCs in various serum-free media has also been tested on the microcarrier platform. Adaptation of hESCs to serumfree medium is usually carried out in organ culture dishes prior to expanding in microcarrier cultures. We have observed that HES-3 on mTeSR1 (Stemcell Technologies) or STEMPRO hESC SFM (Invitrogen) exhibited different growth rates on cellulose microcarriers compared to their counterparts on tissue culture plates, whereas in conditioned hESC medium these cells had similar doubling time for both microcarrier and two-dimensional culture. Microcarrier cultures achieve higher

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cell densities than two-dimensional cultures, and cells in both platforms have a stable karyotype. In addition, we have also successfully cultured two human induced pluripotent stem cells on this platform.

Anticipated Results This method of expanding hESCs on microcarriers using static 6-well plates containing 4 mg/ml of Matrigel-coated cellulose microcarriers per well (working volume of 5 ml per well) is expected to generate about 1.1 ± 0.3 × 106 cells/ml, after 7 days, with an initial seeding density of 0.8 × 105 cells/ml. In a 100-ml spinner ßask, we have observed a cell concentration of up to 3.5 × 106 cells/ml at day 5. For generation of 109 cells, a culture volume of 290 ml is needed. Growing cells for more than 7 days results in low pH and high lactate levels, which lead to arrest in cell growth. We believe higher cell densities can be achieved in a bioreactor controlled for pH and lactate levels.

Time Considerations Preparation of the pre-coated bottles/spinner ßask, thawing Matrigel, and preparing conditioned hESC growth medium is usually started 1 day in advance prior to coating microcarriers. The minimum time required to prepare and coat microcarriers is ∼9 hr. Once the cells have seeded on microcarriers, it should take at least 5 weeks (5 passages) to conÞrm the stability of the culture. Theoretically, starting from one well of a static microcarrier culture in a 6-well plate, ten wells can be seeded in the second week (at about 1:10 split ratio), followed by inoculation of these wells into the bioreactor by the third week. Theoretically, these cells can be seeded into a 1.6-liter bioreactor at an initial cell density of 2 × 105 cells/ml, achieving at least 3.2 × 109 cells by day 5. For a 100-ml spinner ßask with 50-ml working volume, three wells will usually sufÞce, which would only take 1 week for seeding from one well.

Acknowledgement This work is generously supported by Agency for Science, Technology and Research (A*STAR), Singapore.

Literature Cited Altman, S.A., Randers, L., and Rao, G. 2008. Comparison of trypan blue dye exclusion and ßuorometric assays for mammalian cell viability determinations. Biotechnol. Prog. 9:671-674.

Bajpai, R., Lesperance, J., Kim, M., and Terskikh, A.V. 2008. EfÞcient propagation of single cells: Accutase-dissociated human embryonic stem cells. Mol. Reprod. Dev. 75:818–827. Baker, D.E.C., Harrison, N.J., Maltby, E., Smith, K., Moore, H.D., Shaw, P.J., Heath, P.R., Holden, H., and Andrews, P.W. 2007. Adaptation to culture of human embryonic stem cells and oncogenesis in vivo. Nat. Biotechnol. 25:207-215. Braam, S.R., Zeinstra, L., Litjens, S., Ward-van Oostwaard, D., van den Brink, S., van Laake, L., Lebrin, F., Kats, P., Hochstenbach, R., Passier, R., Sonnenberg, A., and Mummery, C.L. 2008. Recombinant vitronectin is a functionally deÞned substrate that supports human embryonic stem cell self-renewal via alphavbeta5 integrin. Stem Cells 26:2257-2265. Choo, A.B.H., Padmanabhan, J., Chin, A.C.P., and Oh, S.K.W. 2004. Expansion of pluripotent human embryonic stem cells on human feeders. Biotechnol. Bioeng. 88:321-331. Choo, A., Padmanabhan, J., Chin, A., Fong, W.J., and Oh, S.K.W. 2006. Immortalized feeders for the scale-up of human embryonic stem cells in feeder and feeder-free conditions. J. Biotechnol. 122:130-141. Ellerstr¨om, C., Strehl, R., Noaksson, K., Hyllner, J., and Semb, H. 2007. Facilitated expansion of human embryonic stem cells by single-cell enzymatic dissociation. Stem Cells 25:1690-1696. GE Healthcare. Microcarrier Cell Culture: Principles and Methods. Handbook. Hasegawa, K., Fujioka, T., Nakamura, Y., Nakatsuji, N., and Suemori, H. 2006. A method for the selection of human embryonic stem cell sublines with high replating efÞciency after single-cell dissociation. Stem Cells 24:26492660. Hu, W.S. and Wang, D.I.C. 1987. Selection of microcarrier diameter for the cultivation of mammalian cells on microcarriers. Biotechnol. Bioeng. 30:548-557. Lin, A.A., Nguyen, T., and Miller, W.M. 1991. A rapid method for counting cell nuclei using a particle sizer/counter. Biotechnol. Tech. 5:153156. Lock, L.T. and Tzanakakis, E.S. 2009. Expansion and differentiation of human embryonic stem cells to endoderm progeny in a microcarrier stirred-suspension culture. Tissue Eng. Part A 15:2051-2063. Malda, J. and Frondoza, C.G. 2006. Microcarriers in the engineering of cartilage and bone. Trends Biotechnol. 24:299-304. Mummery, C.L. 2005. Cardiology: Solace for the broken-hearted? Nature 433:585-587. Nie, Y., Bergendahl, V., Hei, D.J., Jones, J.M., and Palecek, S.P. 2009. Scalable culture and cryopreservation of human embryonic stem cells on microcarriers. Biotechnol. Prog. 25:20-31. Oh, S.K.W., Chen, A.K., Mok, Y., Chen, X., Lim, U.-M., Chin, A., Choo, A.B.H., and Reveny, S.

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2009. Long-term microcarrier suspension cultures of human embryonic stem cells. Stem Cell Res. 2:219-230.

Skottman, H. and Hovatta, O. 2006. Culture conditions for human embryonic stem cells. Reproduction 132:691-698.

Phillips, B.W., Horne, R., Lay, T.S., Rust, W.L., Teck, T.T., and Crook, J.M. 2008. Attachment and growth of human embryonic stem cells on microcarriers. J. Biotechnol. 138:24-32.

Thomas, R.J., Anderson, D., Chandra, A., Smith, N.M., Young, L.E., Williams, D., and Denning, C. 2009. Automated, scalable culture of human embryonic stem cells in feeder-free conditions. Biotechnol. Bioeng. 102:636-1644.

Reubinoff, B.E., Pera, M.F., Fong, C.-Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: somatic differentiation in vitro. Nat. Biotechnol. 18:399404. Richards, M., Fong, C.-Y., Chan, W.-K., Wong, P.-C., and Bongso, A. 2002. Human feeders support prolonged undifferentiated growth of human inner cell masses and embryonic stem cells. Nat. Biotechnol. 20:933-936. Sandford, K.K., Earle, W.R., Evans, V.J., Waltz, H.K., and Shannon, J.E. 1951. The measurement of proliferation in tissue cultures by enumeration of cell nuclei. J. Natl. Cancer Inst. 11:773795. Shah, D., Naciri, M., Clee, P., and Al-Rubeai, M. 2006. NucleoCounter: An efÞcient technique for the determination of cell number and viability in animal cell culture processes. Cytotechnology 51:39-44.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts Science 282:1145-1147. Watanabe, K., Ueno, M., Kamiya, D., Nishiyama, A., Matsumura, M., Wataya, T., Takahashi, J.B., Nishikawa, S., Nishikawa, S.-i., Muguruma, K., and Sasai, Y. 2007. A ROCK inhibitor permits survival of dissociated human embryonic stem cells. Nat. Biotechnol. 25:681-686. White, L.A. and Ades, E.W. 1990. Growth of Vero E-6 cells on microcarriers in a cell bioreactor. J. Clin. Microbiol. 28:283-286. Xu, C., Inokuma, M.S., Denham, J., Golds, K., Kundu, P., Gold, J.D., and Carpenter, M.K. 2001. Feeder-free growth of undifferentiated human embryonic stem cells. Nat. Biotechnol. 19:971-974.

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Germ Layer Induction in ESC—Following the Vertebrate Roadmap

UNIT 1D.1

Jim Smith,1 Fiona Wardle,1 Matt Loose,2 Ed Stanley,3 and Roger Patient4 1

Wellcome Trust/Cancer Research UK Gurdon Institute, University of Cambridge, Cambridge, United Kingdom 2 Institute of Genetics, University of Nottingham, Nottingham, United Kingdom 3 Monash Immunology and Stem Cell Laboratories, Monash University, Clayton, Victoria, Australia 4 Weatherall Institute of Molecular Medicine, University of Oxford, Oxford, United Kingdom ABSTRACT Controlled differentiation of pluripotential cells takes place routinely and with great success in developing vertebrate embryos. It therefore makes sense to take note of how this is achieved and use this knowledge to control the differentiation of embryonic stem cells (ESCs). An added advantage is that the differentiated cells resulting from this process in embryos have proven functionality and longevity. This unit reviews what is known about the embryonic signals that drive differentiation in one of the most informative of the vertebrate animal models of development, the amphibian Xenopus laevis. It summarizes their identities and the extent to which their activities are dose-dependent. The unit details what is known about the transcription factor responses to these signals, describing the networks of interactions that they generate. It then discusses the target genes of these transcription factors, the effectors of the differentiated state. Finally, how these same developmental programs operate during germ layer formation in the context of ESC C 2007 by John differentiation is summarized. Curr. Protoc. Stem Cell Biol. 1:1D.1.1-1D.1.22.
Wiley & Sons, Inc. Keywords: embryonic signals r transcription factors r differentiation r Xenopus r zebrafish r mouse r mesendoderm r mesoderm r endoderm r ectoderm r neural r Wnt r TGF-β r nodal r activin r BMP r FGF r antagonists r T-box r VegT r brachyury r Mix r Sox r GATA r ES cells

LESSONS FROM FROGS (AND FISH) Inductive Interactions in the Early Embryo The most important mechanism by which cell fate is determined in the developing vertebrate embryo involves inductive interactions, in which one group of cells makes a signal that is received, and acted upon, by adjacent tissue. Interactions of this sort are most easily studied in embryos that are accessible to the experimenter, including those of the chick, zebrafish, and frog, and among these species, most insights have come from the frog, Xenopus laevis. In this introductory unit the authors summarize these results, discuss how they might apply to other species, including mammalian embryos, and then consider to what extent they might allow the experimenter to manipulate the differentiation of embryonic stem cells

(ESC). The unit focuses on early developmental decisions, in which cells become committed first to endoderm, mesoderm, or ectoderm, and then to particular regions of these germ layers, such as dorsal versus ventral mesoderm or neural versus non-neural ectoderm. This is done because it is necessary that cultured ESC successfully negotiate these early decisions before they follow pathways leading to the formation of specialized cell types such as pancreas, heart, or liver.

The first step: Dorsalization of the embryo Most of the understanding on inductive interactions during early vertebrate development derives from work on the amphibian species, Xenopus laevis. Xenopus offers many advantages to the developmental biologist: the embryo is large, and therefore easy to inject,

Current Protocols in Stem Cell Biology 1D.1.1-1D.1.22 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01d01s1 C 2007 John Wiley & Sons, Inc. Copyright


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dissect, and manipulate; it is available in large numbers, and therefore suitable for biochemical approaches; it has, at early stages, a reliable fate map, so that one can reliably interpret experiments that are designed to alter cell fates; and its cells are laden with yolk, so that they can survive in a very simple salts solution in the absence of poorly characterized components such as fetal bovine serum. Together, these advantages have made Xenopus the organism nonpareil for the analysis of inductive interactions. When it is laid, the Xenopus egg is a sphere about 1.4-mm in diameter. It has a heavily pigmented animal hemisphere, which comes to lie uppermost in the water, and a paler, yolkier, vegetal half that forms the southern hemisphere. At this early stage, and, as far as is known, in contrast to the mammalian egg, some RNAs are differentially localized within this large cell. Two such RNAs are of particular note. One encodes VegT (also known as Brat, Xombi, and Antipodean), a member of the T-box family of transcription factors (Lustig et al., 1996; Stennard et al., 1996; Zhang and King, 1996; Horb and Thomsen, 1997), and the other encodes Vg1, a member of the transforming growth factor type β (TGF-β) family (Weeks and Melton, 1987). More of these will be discussed later in the unit. For now, suffice it to say that these RNAs move to their vegetal positions during oogenesis, and that their position in this region of the embryo is crucial for normal development. Immediately after being laid the egg appears, and is, radially symmetrical about the animal-vegetal axis. Polarity is imposed upon the egg at the time of fertilization, when the position of sperm entry defines the side of the egg that will eventually form posterior and ventral structures (Vincent and Gerhart, 1987). The mechanism by which this occurs is not well understood, but it is associated with the rotation, of ∼30◦ , of a shell of cortical cytoplasm, just beneath the egg plasma membrane (Vincent and Gerhart, 1987). The position of sperm entry defines the orientation of this rotation, and the significance of the rotation for the specification of the anterior/dorsal to posterior/ventral axis is demonstrated by experiments in which the rotation is inhibited. For example, this can be done by irradiating the vegetal hemisphere of the newly fertilized embryo with UV light (Scharf and Gerhart, 1980; Holwill et al., 1987). This prevents rotation and the embryo develops without a head or axial structures. All that is formed is a mass of ventral tissue that includes, for example,

large amounts of blood (Cooke and Smith, 1987). The loss of anterior tissue is not due to nonspecific effects of UV irradiation, because rotation can be restored by tipping the fertilized egg to one side, and this completely rescues the embryo, such that it forms a perfectly normal tadpole (Scharf and Gerhart, 1980). It is not known how rotation specifies dorsal and anterior regions of the embryo, although it is clear that signaling by members of the Wnt pathway is involved. Thus, ectopic expression of members of the Wnt family in the early Xenopus embryo causes the formation of an additional head (McMahon and Moon, 1989; Smith and Harland, 1991; Sokol et al., 1991), and inhibition or stimulation of Wnt signaling causes, respectively, the loss or enhancement of anterior/dorsal structures (Heasman et al., 1994; Kofron et al., 2001). Most significantly, depletion of maternal Wnt11 mRNA causes embryos to develop without a head or anterior structures, a phenotype that can be rescued by the re-introduction of exogenous Wnt11 mRNA (Tao et al., 2005). Rotation of the cortical layer of cytoplasm is now thought to reposition maternal Wnt11 mRNA from its original vegetal location to a position closer to the equator of the embryo, where it is free to diffuse into deeper cytoplasm. This allows Wnt11 protein to accumulate in dorsal vegetal cells and to activate the canonical Wnt signal transduction pathway in this region of the embryo. It is also possible that cortical rotation causes the dorsal enrichment of Wnt co-receptors, or components of the Wnt signal transduction pathway, in a mechanism that would further enhance the level of Wnt signaling in this region of the embryo (Dominguez and Green, 2000). One of the consequences of Wnt signaling in the dorso-vegetal region of the embryo is the activation of the Siamois gene, which is involved in the formation of dorsal and axial tissues (Lemaire et al., 1995; Fan and Sokol, 1997; Engleka and Kessler, 2001).

The second step: Mesoderm induction As is remarked above, there is no evidence yet for localized RNAs or other determinants in the mammalian egg, and indeed the extent to which the early mammalian embryo is patterned at preblastocyst stages remains controversial (Hiiragi et al., 2006; ZernickaGoetz, 2006). However, at later stages, similarities between amphibian and amniote development are more obvious, and the analysis of mesoderm induction in Xenopus has indeed

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informed studies of mouse development and of ESC differentiation (Gadue et al., 2006). The mesoderm of the amphibian embryo derives from the equatorial region of the embryo. Pioneering experiments by Nieuwkoop showed that this germ layer (and part of the endoderm) forms in this position as the result of an inductive interaction in which cells of the vegetal hemisphere (which inherit, for example, Vg1 RNA and protein from the vegetal region of the fertilized egg) act on overlying cells (Nieuwkoop, 1969; Sudarwati and Nieuwkoop, 1971). Nieuwkoop demonstrated this interaction by dissecting cells from the animal pole region of the embryo (which normally become ectoderm), and juxtaposing them with vegetal cells (which normally become endoderm). Individually neither cell population would form mesoderm, but the combination of cells produced large amounts of mesodermal tissue such as muscle, and lineage labeling experiments revealed that the mesoderm derived from the ectodermal tissue, indicating that the inducing signal derives from the endoderm. Mesoderm inducing factors Work in the late 1980s and early 1990s revealed that two classes of signaling molecules have the ability to mimic the effect of the vegetal cells. These include members of the fibroblast growth factor (FGF; Slack et al., 1987; Kimelman et al., 1988) and transforming growth factor type β (TGF-β) families (Albano et al., 1990; Asashima et al., 1990; Smith et al., 1990; Thomsen et al., 1990). The most powerful inducers proved to be members of the TGF-β family, and especially activin and the Xenopus nodal-related (Xnr) genes (Jones et al., 1995; Joseph and Melton, 1997; Takahashi et al., 2000), and since this time two other related proteins have been characterized as mesoderm-inducing factors: Vg1, whose transcripts are expressed in the vegetal region of the oocyte and egg (Weeks and Melton, 1987; Dale et al., 1993; Thomsen and Melton, 1993; Birsoy et al., 2006), and derri`ere, which, like activin and the nodal-related genes, is expressed zygotically (Sun et al., 1999a). When applied to isolated animal pole regions, all of these inducing factors, including members of both the FGF and TGF-β families, can cause the activation of mesoderm- and endoderm-specific genes and the differentiation of mesodermal cell types. One difference between the two families of signaling molecules is that FGF family members induce predominantly ventral and posterior

cell types, and fail to induce the expression of genes that are expressed in prospective anterior and dorsal tissues, such as goosecoid (Green et al., 1990). In contrast, the members of the TGF-β family tend to induce a wide spectrum of tissues, from ventral and posterior to dorsal and anterior, and they do this in a concentration-dependent manner, with lower concentrations inducing posterior/ventral tissues and high doses inducing anterior and dorsal structures (Green and Smith, 1990; Green et al., 1992; Green, 1994; Gurdon et al., 1994, 1995, 1996). These dose-dependent effects of the TGF-β family are discussed below. The FGF and the TGF-β families employ different signal transduction pathways, the former inducing mesoderm through the MAP kinase pathway (Gotoh et al., 1995; LaBonne et al., 1995; Umbhauer et al., 1995) and the latter through the Smad family (Hill, 2001). Limited knowledge of the transcriptional regulation of known FGF and TGF-β target genes (Watabe et al., 1995; Howell and Hill, 1997; Latinkic et al., 1997; Howell et al., 1999; Germain et al., 2000; Lerchner et al., 2000) provides a molecular basis for the difference in the inducing activities of the two classes of signaling molecules. Most significant, however, is the conclusion confirmed by one of the first experiments to employ transgenesis in Xenopus species that the role of FGF signaling is to maintain mesodermal identity rather than to induce it (Kroll and Amaya, 1996). Concentration-dependent effects of inducing factors It is interesting that members of the TGFβ family are able to induce different types of mesoderm, and to activate the expression of different mesoderm- and endoderm-specific genes, at different concentrations. Unfortunately, rather little is known about the mechanism through which this occurs. Work by Gurdon and colleagues has shown that 100 molecules of activin bound to a single animal pole cell are sufficient to induce expression of the pan-mesodermal gene brachyury, while 300 molecules are required to extinguish brachyury and to activate goosecoid (Dyson and Gurdon, 1998). It is also known that protein synthesis is required for the downregulation of brachyury expression that occurs at high doses of activin, suggesting that these high doses of a TGF-β family member induce a repressor of brachyury (Papin and Smith, 2000). One candidate for such a repressor is Goosecoid itself, and indeed mutation of a Goosecoid binding site in a 381-base

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pair brachyury promoter fragment prevents the down-regulation of a brachyury reporter construct that occurs at high activin concentrations (Latinkic et al., 1997). However, it is unlikely that Goosecoid is the only repressor of brachyury that is induced at high levels of activin, because inhibition of Goosecoid activity does not affect the down-regulation of endogenous brachyury that occurs at high activin concentrations (Papin and Smith, 2000). This phenomenon requires further investigation, because, as is discussed below, it is likely that pattern formation in the developing embryo occurs in response to gradients of TGF-β family members, and attempts to direct ESC differentiation must take account of the concentrations of the inducing factors that are used. The most direct evidence that gradients of inducing factors activate different genes in different regions of the embryo, and specify different cell types, comes from experiments in which the functions of the genes are inhibited. Developmental biologists know of eight TGFβ family members that are expressed in the embryo and that have mesoderm-inducing activity: Vg1, activin, Xnr1, Xnr2, Xnr4, Xnr5, Xnr6, and derri`ere (see references above). It has not yet proved possible to inhibit the action of all of these factors individually, but antisense and dominant-negative approaches have demonstrated that the maternally expressed gene Vg1 (Birsoy et al., 2006), and the zygotically activated activin (Piepenburg et al., 2004) and derri`ere (Sun et al., 1999a) are all required for proper mesoderm formation. Of the nodalrelated genes, only Xnr1 has been studied individually (Toyoizumi et al., 2005), and it proves to be required for proper specification of the left-right axis of the embryo. However, simultaneous inhibition of all the Xenopus nodalrelated genes by increasing expression of a truncated form of Cerberus, termed Cerberusshort, causes the progressive loss of dorsal and then ventral gene expression, consistent with the idea that gradients of these TGF-β family members specify cell types in the developing embryo (Agius et al., 2000). It is not yet clear how such gradients are established or how the inducing factors traverse fields of responding cells, although the higher expression of the nodal-related genes at the dorsal side of the embryo is thought to derive from the enhanced VegT, Vg1, and Wnt signaling in this region of the embryo (Agius et al., 2000), and it seems likely that inducing factors travel in the extracellular milieu rather than through an intracellular

route such as transcytosis (Williams et al., 2004).

Refining the pattern: Inhibiting BMP signaling Different concentrations of TGF-β family members, such as activin and the nodalrelated genes, can establish differences between the prospective anterior/dorsal and posterior/ventral regions of the late blastula and early gastrula regions of the embryo. These differences, however, are rather crude. Dissection of tissue from different regions along this axis of the embryo reveals that there are only two well-defined domains, a smaller one on the dorsal side of the embryo, where the dorsal lip of the blastopore will appear, and a larger one that comprises the rest of the equatorial region of the embryo (Dale et al., 1985). Culture of the former region reveals that this is specified to form notochord and muscle, while the latter region forms predominantly blood. Significantly, these results stand in contrast to the fate map of the embryo at this stage, which shows that most of the muscle of the embryo derives from the larger, ventral, region of the prospective mesoderm (Dale and Slack, 1987). How can these observations be reconciled? Further grafting experiments show that the region of the embryo that will form the dorsal lip of the blastopore is the source of signals that can dorsalize adjacent ventral mesoderm (Smith and Slack, 1983). Indeed, if this region, known as Spemann’s organizer, is grafted to the ventral region of a host embryo, the host develops a secondary axis on its ventral side (Spemann, 1938). Interactions between Spemann’s organizer and the rest of the embryo refine the spatial patterns of gene expression and cell differentiation along the anterior/dorsal to posterior/ventral axis. Identification of the signals produced by Spemann’s organizer that are responsible for dorsalization came from several types of experiments. Expression cloning identified the gene noggin (Smith and Harland, 1992), which was capable of rescuing embryos that had been ventralized by treatment with UV light (see above). An in situ hybridization screen showed that chordin is expressed in the dorsal region of the embryo, with subsequent experiments revealing that it too had dorsalizing activity (Sasai et al., 1994); and analysis of follistatin, whose gene product was already known to inhibit activin signaling, showed that this gene is also expressed dorsally and is capable of dorsalizing ventral mesoderm (Hemmati-Brivanlou et al., 1994). All of these factors have in common the fact that they can

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inhibit signaling by bone morphogenetic proteins (BMPs), factors which previously had been shown to have powerful ventralizing activity in the Xenopus embryo (Dale et al., 1992; Jones et al., 1992), and subsequent work demonstrated that these inhibitors can set up a gradient in the embryo, high in the anterior/dorsal region and low in posterior/ventral tissues (Jones and Smith, 1998). BMP family members such as BMP4 are expressed in a widespread fashion throughout the embryo (with the exception of the dorsal region; Hemmati-Brivanlou and Thomsen, 1995), so that the gradient of BMP inhibitors establishes a reverse gradient of BMP activity. It is this gradient that creates positional information along the anterior/dorsal to posterior/ventral axis of the embryo. Like Vg1, activin, the Xnr proteins, and derri`ere, BMPs are members of the TGF-β family, but they differ because they signal through different cell surface receptors and different members of the Smad family. Activin and the other mesoderm-inducing factors signal through Smad2 or Smad3, which form heteromeric complexes with Smad4; BMPs signal through Smad1, Smad5, and Smad8, which also form complexes with Smad4 (Hill, 2001; Schier, 2003).

The organizer produces many inhibitors The discovery of noggin, chordin, and follistatin as factors that pattern the anterior/dorsal to posterior/ventral axis of the embryo by inhibiting BMP signaling was quickly followed by the identification of other inhibitors of extracellular signaling molecules. These include Frzb-1 (Leyns et al., 1997), a secreted protein containing a domain similar to the putative Wnt-binding region of the frizzled family of transmembrane receptors, and Dickkopf-1 (Dkk1; Glinka et al., 1998), a ligand for the Wnt co-receptor LRP6. Both of these molecules inhibit signaling by Wnt family members, and have the effect of inhibiting the ventralizing effects of proteins such as Wnt8, which is expressed throughout the lateral and posterior/ventral regions of the early gastrula-staged embryo. They therefore establish an effective reverse gradient of Wnt signaling, further refining positional information along the dorso-ventral axis of the embryo. Perhaps the most remarkable inhibitor produced by Spemann’s organizer, however, is Cerberus, which inhibits the activities of BMP, Wnt, and nodal-related signaling (Bouwmeester et al., 1996; Glinka et al., 1997;

Piccolo et al., 1999). Ectopic expression of Cerberus in the Xenopus embryo causes the formation of an extra head (Bouwmeester et al., 1996), indicating, remarkably, that formation of the head does not require special or novel inducing activities but rather the inhibition of several signaling pathways.

Later inductive interactions and implications for embryonic stem cells This description of the induction and patterning of the mesoderm has emphasized (1) the importance of intercellular signaling by secreted polypeptide growth factors, (2) the fact that these molecules can exert concentrationdependent effects, and (3) the observation that effective gradients of such growth factors can be established by reverse gradients of secreted inhibitors. These principles also apply to later stages of embryonic development. For example, neural induction by Spemann’s organizer in the gastrula-stage embryo is largely a consequence of the inhibition of BMP signaling in the prospective ectoderm (Harland, 1994), together with contributions from FGF and probably other signaling families (Launay et al., 1996; Sasai et al., 1996; Linker and Stern, 2004). At later stages and in other tissues, other families of signaling molecules also play a role, including hedgehog and notch signaling. A thorough knowledge of these signaling events will be invaluable in coming to understand the ways in which one can direct and influence ESC differentiation.

Integration of Intracellular Responses to Embryonic Induction The distinct combinations and levels of embryonic signals experienced by different cells in the embryo differentially induce the expression or activities of transcription factors (TFs); and the particular combination of TFs active in a given cell determines its phenotype. The responses of TFs to embryonic signals together with interactions between TFs constitute a genetic regulatory network (GRN), describing the state of a given cell at a particular time (Loose and Patient, 2004; Koide et al., 2005). To understand germ layer induction, it is necessary to fully understand the construction, dynamics, and stability of these networks as differentiation proceeds. Here, because of the volume of data available, the authors concentrate on mesendoderm induction in Xenopus, but much of what has been found in Xenopus is the same or very similar in zebrafish, making it very likely that the fundamental mechanisms discussed here

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Legend at right.

serve as a model for vertebrate germ layer formation generally. In vertebrates, the three primary germ layers, ectoderm, mesoderm, and endoderm, are classically regarded as the first signs of differentiation in the embryo proper, with differentiation to extra-embryonic tissues taking place concomitantly. However, a number of observations suggest that the initial separations may not be as clean as previously thought (Rodaway and Patient, 2001; Wardle and Smith, 2004). In addition, subdivisions within the mesoderm become apparent in some cases before mesodermal and endodermal fates have been distinguished. Thus, skeletal muscle and notochord progenitors become distinguishable from endodermal precursors before blood and cardiovascular progenitors. Observations of this nature have led to the use of the term mesendoderm, and comparisons with more primitive organisms indicate that the distinction may in fact be quite ancient.

In order to begin to appreciate the implications of the Xenopus network for our understanding of ESC programming, the authors have built a simplified version of this network, which takes account of reduced gene numbers in mammals and illustrates the key principles (Fig. 1D.1.1A). GRNs are composed of many smaller networks, termed motifs (for a review see Lee et al., 2002 or Babu et al., 2004). These motifs occur more often than would be expected in a random network with the same degree of connectivity as the network under study. Such motifs are easiest to identify computationally, and the program mFinder has been used to identify motifs in a recent update of the Xenopus mesendoderm network used (Milo et al., 2002; Loose and Patient, 2004). The complete network is scale-free, that is, all genes do not display equal connectivity. Some of the TFs studied to date display substantially more connectivity than others making them potentially master regulators. The following

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will highlight some of these and the motifs found in the Xenopus GRN, and will try to draw out the lessons to be learned for ESC differentiation.

Feed forward loops The most striking thing about mesendoderm formation in Xenopus is the extent to which the maternal T-box TF, VegT, dominates proceedings (Zhang et al., 1998; Xanthos et al., 2001; Taverner et al., 2005; Heasman, 2006). Of 54 genes currently in the network, 26 are regulated directly by VegT. A clue to its function in the network emerges when considering which network motifs feature VegT. VegT features heavily in a common motif, the feed forward loop (see Fig. 1D.1.1B). Feed forward loops consist of three genes, X, Y, and Z, where X regulates Y and Z, and Y regulates Z (Milo et al., 2002). Studies of the properties of feed forward loops reveal two likely functions: either accelerated or delayed expression of the target Z, depending on the nature of the interactions between the three genes (activating or repressing) and the way in which the X and Y genes interact at Z (both or either required for expression; see Mangan et al., 2003). Of the 63 such feed forward motifs identified in the Xenopus network, 41 are initiated by VegT. Furthermore, the majority of these 41 motifs

are of the form that would be expected to accelerate the onset of expression of the target. The targets of the feed forward motifs in the Xenopus GRN include both mesodermal and endodermal TFs, suggesting that cells in which maternal VegT is expressed have the potential to become both mesoderm and endoderm, i.e., a molecular demonstration of their mesendodermal potential. Of the remaining 22 feed forward loops identified in the updated Xenopus mesendodermal network, 13 are initiated by nodal-related signals, 5 by β-catenin, driven maternally by Wnt11 signaling (Tao et al., 2005), and 4 by Mix family TFs or GATA factors. Thus, in the Xenopus embryo, the localization of maternal determinants is a crucial aspect of mesendoderm formation. The significance of this for programming ESCs remains to be seen. For example, even though zebrafish express a maternal T-box TF, its activities have not yet been demonstrated to be equivalent to VegT in Xenopus (Bjornson et al., 2005), so it is possible that this is a derived characteristic of Xenopus. Alternatively, the loss of such a wide range of activities might be a derived characteristic of zebrafish, in which case, one should be looking for the distribution of T-box transcription factors in ESCs and determining the extent to which they

Figure 1D.1.1 (at left) (A) A reduced network representing the key interactions occurring within the Xenopus mesendoderm network at the start of gastrulation (approximately stage 10.5; Loose and Patient, 2004). In order to reduce the complexity of the network, the authors have combined the interactions of key gene family members into one representative gene. Thus, the nodal family members (and derriere) are combined into one nodal gene, the mix family members into one mix ` and the GATA genes into one GATA gene. For the nodal and mix families, this reduction reflects the fact that there is only one mix and one nodal homolog in mouse and human. In contrast, the reduction of the GATA family into one representative reflects the difficulty of distinguishing the targets of individual GATA factors at this time, although recent work has begun to make some headway (Afouda et al., 2005). The interactions shown in the diagram represent the combined input and output of all family members in the original network. No explicit conflicts were found within families (i.e., one family member positively regulating, another negatively), although for some interactions, individual members have been shown to regulate a subset of the targets of the whole family. In these cases the authors have assumed that the ability to regulate a given target has been lost by the individual family member as opposed to having been gained by several family members. In order to clearly represent the topology of the network, genes that do not feed back into the network have not been shown. The exception to this is the two targets of the GATA family, sox17 and HNF1β. At later time-points these genes will feed back into the network. sox17 and HNF1β also represent endoderm within the network. Dashed lines illustrate interactions that are initiated by a maternal message, although later, zygotic expression will take over. Examples of feed forward loops (B) can be traced in this network, for example VegT to nodal to mix (illustrated in C) and mix to GATA to sox17. (B) Illustration of a feed forward motif. Gene X activates both Y and Z, gene Y activates Z. A feed forward loop can have any combination of activation and repression, resulting in 8 different sub-classes of feed forward loop. (C) A sample feed forward loop involving VegT, nodal, and mix. Note that nodal is able to autoregulate and can therefore maintain mix expression in the absence of VegT. (D) This motif is extracted from panel A and shows the antagonism between mix and brachyury. In addition, goosecoid is activated by mix and in turn repressed brachyury expression.

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correlate with mesendoderm differentiation. In the absence of a maternal T-box TF, expression of a T-box TF could be induced by FGF as seen for Brachyury and FGF4 (eFGF) in the Xenopus mesendoderm network (Isaacs et al., 1994; Casey et al., 1998; see below). Thus, in the absence of a maternal T-box transcription factor with the full range of activities in ESCs, FGF signaling may play a greater role in establishing the mesoderm and the endoderm. As discussed below, nodal-related signaling is often downstream of VegT and the nodal-related family members share many of the targets of VegT. Therefore, another possible substitute for a maternal T-box determinant could be nodal signaling itself. Finally, in the case of maternal β-catenin, whose nuclear localization is driven by maternal Wnt11, the evidence suggests that this pathway is intact in ESCs (Lindsley et al., 2006).

Autoregulation An important consideration when driving a cell along a specific differentiation pathway is how the cell will maintain the new pathway once the driving stimulus has been removed. In addition to the example of FGF and Brachyury mentioned above, the key initial driver of the Xenopus mesendoderm network, VegT, drives expression of the nodals, of which Xnr1, 2, and 4 can positively autoregulate (Osada et al., 2000; Cha et al., 2006). Of the 63 feed-forward motifs identified thus far, 27 involve a nodal family member in the Y position of the motif (see Fig. 1D.1.1B,C). Thus, the initial activating push that is provided by VegT is gradually replaced by nodal signaling, and this signal has the ability to maintain its expression and the pathway remains active as VegT is depleted. Thus, autoregulation is another key motif in genetic regulatory networks contributing to cell memory and forward momentum. Buffering of positive autoregulation is provided by negative regulators, such as antivin/lefty, itself a target of nodal signaling (Cha et al., 2006).

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An important concept in differentiation, in addition to activation of specific programs, is the shutting down of alternative pathways. A control motif unique to cells that have to make lineage decisions is cross-antagonism. In the most complete examples, this motif has two master regulators each trying to drive different programs, with each regulator positively autoregulating its own expression, providing a driving force for that particular program. In

addition to this, each factor antagonizes the other’s activities and so inactivates the alternative pathway. Examples of this can be found in lineage decisions in the blood. Here, GATA1 drives the erythroid/megakaryocyte program and the Ets factor, Pu.1, drives other myeloid outcomes (Nerlov et al., 2000; Zhang et al., 2000; Swiers et al., 2006). This switch can be modified by altering the levels of the master regulators (Galloway et al., 2005; Rhodes et al., 2005). The closest motif with characteristics of cross-antagonism in the Xenopus mesendoderm network involves Mix.1 and Brachyury. These TFs repress each other’s expression and likely in part underpin the choice between endoderm and mesoderm from the mesendodermal layer (Lemaire et al., 1998). Although there is no evidence for direct autoregulation of Mix.1 or Brachyury, the autoregulatory loop between Brachyury and FGF4 provides this function. The apparent absence of autoregulation on the Mix.1/endoderm pathway may be compensated for by the role of Goosecoid, which throws the switch towards Mix.1/endoderm, because it represses Brachyury and is itself downstream of Mix.1 (see Fig. 1D.1.1D; Latinkic et al., 1997; Germain et al., 2000). This relationship between Goosecoid and Brachyury likely also underpins the anteroposterior patterning of the mesoderm, with continued Brachyury expression being restricted to the posterior mesoderm by Goosecoid repression. Mix.1 favors endoderm differentiation by driving expression of GATA factors and Sox17. Both factors drive expression of endodermal genes throughout the future endoderm in Xenopus and zebrafish, and, later in the anterior of the embryo. Sox17 also regulates Goosecoid positively, thereby reinforcing the endodermal lineage decision there (Latinkic and Smith, 1999; Sinner et al., 2004, 2006; see below). Both Mix.1 and Brachyury are downstream of the same nodal-related signaling factors and are initially found co-expressing in the same cells (Lemaire et al., 1998), suggesting that cells are initially programmed with the potential to differentiate into either germ layer and subsequent downstream interactions select one germ layer or another. As already discussed, the choice of germ layer is determined by the combination or concentration of the inducing signals. Thus, individual cells within the mesendoderm may have indistinguishable molecular phenotypes reflecting the broad potential of these cells.

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Network states Microarray based profiling of gene expression in embryos is revealing new interactions important for endoderm and mesoderm specification, and these will need to be incorporated into the GRN (Dickinson et al., 2006; Sinner et al., 2006). However, it is already clear that GRNs are four dimensional, with levels of individual transcription factors varying in both space and time between, and possibly also within, fields of cells. In vivo the network of an individual differentiating cell will pass through various intermediate states and there may be a limited number of ways of constructing the stable network responsible for the phenotype of a specific cell type. A better understanding of these phenomena in developing embryos will likely enable enhanced control over ESC differentiation.

Direct Target Genes The activity of early regulatory transcription factors must be translated into downstream responses, such as the cell movements of gastrulation and the onset of cell differentiation. This is achieved through regulating the expression of downstream targets, which may themselves be regulatory factors or signals giving rise to GRNs as discussed above, or may be direct effectors of cell movement or differentiation. Below the authors discuss some direct targets and the roles they play in mediating the activities of a few of these early regulatory transcription factors in mesendoderm.

Mesoderm Direct targets of Brachyury Brachyury is expressed throughout the mesoderm of early vertebrate embryos as an immediate response to mesoderm induction, and it is required for many aspects of mesoderm formation. Inhibition of Brachyury function and mutations in the gene reveal that its orthologs in mouse, frog, and zebrafish are required during gastrulation for normal morphogenetic cell movements, for notochord and posterior mesoderm formation, and for the establishment of left-right asymmetry (Smith, 2001; Amack and Yost, 2004). Moreover, ectopic activation of Xenopus Brachyury in Xenopus ectodermal explants (animal caps) is sufficient to induce ventral and lateral mesodermal cell fates in those cells (Cunliffe and Smith, 1992). Double mutants uncover additional roles for Brachyury that are not evident in the single mutant due to redundancy with other factors. For instance double mutants in

ntl (no tail, the zebrafish Brachyury homolog) and spt (tbx16) or zygotic oep reveal a role for Brachyury in trunk somite and blood formation in zebrafish (Schier et al., 1997; Amacher et al., 2002). Brachyury exerts its effects on mesodermal cells both through regulating cell fate, as shown by down regulation of target genes when Brachyury activity is decreased or absent, and through regulating cell movements during gastrulation, because cells do not move to their correct positions in brachyury mutant embryos. Research over the last decade or so has also identified a handful of direct targets of Brachyury in Xenopus, including Xwnt11, Vent2b, and FGF4 that mediate both cell fate choices and cell movement (Casey et al., 1998; Tada et al., 1998; Tada and Smith, 2000; Messenger et al., 2005). For instance, Wnt11, acting through the noncanonical Wnt pathway, regulates the convergent extension movements of gastrulation in both Xenopus and zebrafish (Heisenberg et al., 2000; Tada and Smith, 2000). Vent2b, on the other hand, is a transcriptional repressor that plays a role in BMP-mediated specification of ventral and paraxial mesoderm, at least in part through its ability to repress goosecoid expression (Ladher et al., 1996; Onichtchouk et al., 1996; Schmidt et al., 1996; Trindade et al., 1999; Melby et al., 2000). Brachyury binds the promoter of Xvent2b in vivo, and probably does so in combination with Smad1, a transducer of BMP signals, in ventral lateral mesoderm (Messenger et al., 2005). FGFs are involved both in regulating gene expression and cell movement. Inhibition of FGF signaling causes loss of posterior somites and notochord, and defects in gastrulation movements in Xenopus and zebrafish (Amaya et al., 1993; Griffin et al., 1995). Similarly, FGF signaling is required for normal mesoderm formation in mouse embryos (Deng et al., 1994; Yamaguchi et al., 1994). Several different FGFs are expressed in the mesoderm of Xenopus (fgf3, fgf4 [efgf], and fgf8b) and zebrafish (fgf8, fgf17β, and fgf24) and act in combination to regulate gene activity which patterns and maintains mesoderm (Furthauer et al., 1997; Fisher et al., 2002; Draper et al., 2003; Fletcher et al., 2006b). For instance, FGF4 and Brachyury are involved in a positive autoregulatory feedback loop (Isaacs et al., 1994; Casey et al., 1998), and both FGF4 and FGF8b regulate MyoD expression (Fisher et al., 2002; Fletcher et al., 2006b). Inhibition of FGF signaling also causes defects in cell movement during gastrulation, and

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evidence in chick embryos suggests that different FGFs can act as chemoattractants or chemorepellents to ensure that migrating mesodermal cells arrive at their correct destination during gastrulation (Yang et al., 2002). Interestingly, target genes of Brachyury that directly mediate notochord differentiation have yet to be isolated. Furthermore, as mentioned above, during the early stages of mesendoderm formation, Brachyury is expressed in cells that will eventually become endoderm, although expression resolves into mesodermal cells as development proceeds (Rodaway et al., 1999; Wardle and Smith, 2004). Similarly Brachyury-expressing cells in embryoid bodies have the potential to go on and form both mesoderm and endodermal lineages (Fehling et al., 2003; Kubo et al., 2004). Given this, it is possible that Brachyury also regulates the expression of early endodermal genes, although targets are yet to be identified.

Endoderm

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Direct targets of Sox17 In Xenopus, sox17 alleles (there are three) are required in combination for normal endoderm formation, while in zebrafish, Sox17 and the related factor, Sox32/Casanova, both play a role in endoderm formation, with Casanova acting upstream of sox17. Inhibition of all Sox17 activity in Xenopus leads to downregulation of endodermal markers, abnormal gut formation, and inhibition of gastrulation movements (Clements and Woodland, 2000). Similarly, mouse embryos null for sox17 have defects in gut formation (Kanai-Azuma et al., 2002). Direct targets of Sox17 in the gastrula stage Xenopus embryo include endodermin, HNF1β, foxa1, and foxa2 which are expressed in the endoderm of early embryos (Clements and Woodland, 2003; Ahmed et al., 2004; Sinner et al., 2004). foxa2 is also expressed in the floorplate of the neural tube, and mutants in zebrafish foxa2 (monorail) show a defect in floor plate differentiation (Norton et al., 2005). On the other hand, inhibition of Xenopus Foxa2 in ventral tissues using an engrailed repressor construct causes expression of dorsoanterior mesendodermal markers and some decrease in anterior structures, while ectopic expression of foxa2 in Xenopus inhibits the expression of mesoendodermal markers and causes severe defects in gastrulation (Suri et al., 2004). The difference in phenotypes in Xenopus and zebrafish may be due to the engrailed repressor construct inhibiting addi-

tional fox-related factors, the monorail mutation not being a complete null, and/or other factors taking over the role of foxa2 in mutant zebrafish. Inhibiting HNF1β activity in Xenopus using an engrailed repressor construct inhibits mesoderm induction by vegetal explants and causes defects in mesoderm formation if localized to the marginal zone, while inhibition or augmentation of HNF1β activity in Xenopus, using mutated human alleles, leads to defects in pronephros formation (Vignali et al., 2000; Wild et al., 2000). Hence, although these downstream targets play some role in aspects of mesendoderm formation, it is not clear from these experiments which aspects of Sox17 activity they are involved in, and it is evident that more targets remain to be isolated. Targets of GATA factors In Xenopus and zebrafish GATA 4 to 6 mediate endoderm formation. Ectopic expression of GATA 4, 5, or 6 in Xenopus animal caps induces endodermal markers, such as sox17β and HNF1β, while knock down of GATA 5 and 6 activity in the whole embryo using antisense morpholinos leads to defects in gut morphology (Weber et al., 2000; Afouda et al., 2005). In zebrafish, mutations in gata5 (faust) lead to a decrease in early endodermal markers and subsequent defects in gut morphogenesis (Reiter et al., 1999, 2001). Little is known about direct targets of GATA factors, although experiments in which protein synthesis was inhibited suggest that GATA6, and to some extent GATA5, is able to directly activate expression of sox17β and HNF1β in Xenopus (Afouda et al., 2005). GATA factors are also involved in migration of the leading edge mesendoderm across the blastocoel roof during gastrulation (Fletcher et al., 2006a), although the direct targets that mediate this activity are not known. Identification of direct targets Clearly, in order to better understand how the actions of regulatory transcription factors induced in response to mesendoderm-inducing signals are translated into cell movements and the onset of cell differentiation, it is necessary to have a better understanding of the direct targets that mediate those processes, and particularly those targets that directly affect those processes. One method of identifying direct targets is by a candidate approach. For instance, subtractive screens or, more recently, microarray experiments can be used to identify genes

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whose expression is altered in response to activation or inhibition of a transcription factor (Saka et al., 2000; Taverner et al., 2005). Using protein synthesis inhibitors, such as cycloheximide, one can then identify whether these genes are directly regulated transcriptional targets of that factor (Taverner et al., 2005). Another, more high-throughput method of identifying directly bound targets of a particular transcription factor, or other DNA binding protein, in embryos or ESCs is chromatin immunoprecipitation combined with genomic microarrays (Taverner et al., 2004; Boyer et al., 2005; Wardle et al., 2006). In this method, often known as ChIP-chip or location analysis, cells or embryos are treated with formaldehyde to cross-link the proteins to DNA. The DNA is then fragmented (ranging in size from ∼0.2 to 2 kb) and antibodies to the factor of interest are used to immunoprecipitate it and identify the DNA bound to it in vivo. The isolated DNA is then amplified, labeled, and hybridized to a microarray containing genomic promoter sequences. Additional methods to identify genomic sequences directly bound by a factor that have been used in other systems such as Drosophila embryos, C. elegans, or mammalian cell lines include Dam-ID (van Steensel et al., 2001; Greil et al., 2006) and ChIP-cloning (Weinmann et al., 2001; Oh et al., 2006) or ChIP coupled with pair-end ditag sequencing (Wei et al., 2006). Cloning and sequencing the isolated fragments has the advantage of identifying all genomic regions that are associated with the factor, whereas currently genomic microarrays for vertebrate organisms contain only a subset of the genome such as sequences around the gene promoter, although this is likely to change as new technologies allow larger whole genomes to be studied. These powerful direct genomic binding approaches will eventually lead to the identification of all the direct targets of each of the regulatory TFs involved in germ layer formation. These data will enhance the GRNs described above and allow for a more complete understanding of the control circuits of differentiation. They will in addition eventually identify the end game of each differentiation pathway with a complete readout of which regulatory TFs drive each of the terminal differentiation products. Together with the information on the embryonic signals that control the activities of these regulatory TFs, the ability to control differentiation will be substantively improved.

GERM LAYER INDUCTION DURING EMBRYONIC STEM CELL DIFFERENTIATION Developmental Potential of ESCs ESCs are pluripotent immortal cells derived from the inner cell mass of the preimplantation mammalian blastocyst (Evans and Kaufman, 1981; Martin, 1981). The clearest demonstration that ESCs are capable of generating all of the cell types found in the adult comes from experiments in which normal fertile mice composed entirely of ESC-derived cells were generated by aggregating clusters of ESCs with tetraploid embryos (Nagy et al., 1993). In these experiments, instructive signals from the tetraploid embryo–derived extraembryonic tissues initiated a cascade of developmental programs within the ESC aggregates, culminating in the formation of a complete animal. Similarly, the application of exogenous instructive signals to aggregates of ESCs in vitro also initiates a cascade of differentiation resulting in the generation of cells representing multiple tissue types.

Developmental Congruence and Directed Differentiation The factors governing establishment of the embryonic germ layers, ectoderm, mesoderm, and endoderm, have been determined from embryological studies, predominantly using model systems such as zebrafish and Xenopus (Kimelman and Griffin, 2000; Loose and Patient, 2004; Tam et al., 2006). These factors form part of an evolutionarily conserved genetic regulatory network that coordinates gene expression, cell movement, and differentiation. Within these networks, specific genes mark or regulate sequential embryonic stages as cells pass through a series of progressive developmental restrictions culminating in their irreversible commitment to the germ layers, ectoderm, endoderm, and mesoderm. These same steps can be observed during the early phases of ESC differentiation, with cells sequentially expressing genes representing inner cell mass, epiblast, primitive streak, mesoderm, and endoderm (Hirst et al., 2006). Likewise, the commitment of ESC differentiation to a particular developmental program can be precipitated by the same extracellular factors found to be important for the execution of corresponding programs within the embryo (see Fig. 1D.1.2). This correspondence between what happens in the embryo and what happens during ESC differentiation in vitro

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Figure 1D.1.2 Schematic representation of key steps and factors involved in the specification of the germ layers from differentiating ESCs. BMP and Activin-like signals are required for expansion of cell numbers as cells traverse the in vitro equivalent of an epiblast stage (Mishina et al., 1995; Song et al., 1999), marked by expression of FGF5. In the absence of BMP, Wnt, or Nodal signals, epiblast cells adopt a default neurectodermal fate marked by expression of genes including Sox1 or Pax6. In the presence of BMP, Nodal, or Wnt signals, cells commit to the in vitro equivalent of primitive streak formation, marked by expression of genes such as brachyury, Mixl1, and FoxA2. FGF signals may play a role in both the neurectodermal and mesendodermal differentiation pathways (Dell’Era et al., 2003; Sun et al., 1999b; Ying et al., 2003), although this has not yet been assessed in conditions free from extraneous influences. Once mesendoderm formation is initiated, the continued presence of a robust Activin-like signal promotes the formation of anterior-dorsal mesoderm (notochord, somites, cardiac) and definitive endoderm. Whereas, high levels of BMP activity favor the generation of posterior-ventral hematopoietic mesoderm. The overlapping triangles representing Activin, Wnt, and BMP activity serve as a reminder that all of these signaling pathways must be intact for correct patterning of the emerging mesendoderm. Activin* signifies that the identity of the ligand responsible for the activin-like signal at this stage is unclear.

Germ Layer Induction in ESC

represents a developmental congruence between the two systems. The existence of developmental congruence allows studies of embryo development in vivo to be used as a reference for understanding and manipulating ESC differentiation in vitro. A second concept underlying the manipulation of ESC differentiation systems is encapsulated in the phrase “directed differentiation.” This term has been used to describe ESC differentiation protocols that give the ex-

perimenter more of the cell type they desire and a lower proportion of unwanted cell types. Taken together, the ideas of developmental congruence and directed differentiation provide a conceptual framework for the development of ESC differentiation protocols aimed at producing large numbers of specific cell types.

Neural induction A critical idea surrounding the process of germ layer formation is the concept of the

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default differentiation pathway; the commitment of cells to a particular lineage in the absence of signals to the contrary. This idea had its origins in studies of Xenopus development in which it was demonstrated that ectodermal cells deprived of the opportunity for cellcell interactions adopted a neural fate (Grunz and Tacke, 1989; Green, 1994). Subsequent work demonstrated that a blockade of BMP signaling or of the activity of Brachyury, a critical target gene of BMP/Nodal/Wnt signaling, was sufficient to permit neural differentiation of naive ectoderm (Rao, 1994; Hawley et al., 1995; Grunz, 1996; Hansen et al., 1997). These studies were possible because cells of the Xenopus embryo survived in a medium devoid of constituents with inductive activities. Application of this principle applied to ESC differentiation required the development of a medium capable of supporting cell growth and survival without imposing a predetermined differentiation outcome (Johansson and Wiles, 1995; Wiles and Johansson, 1997). Experiments performed using such a chemically defined media (CDM) showed that in the absence of serum, BMPs, or Activin A, differentiating ESCs up-regulated expression of the neural marker Pax6, suggesting cells had embarked on a neural differentiation pathway (Wiles and Johansson, 1997). These and subsequent studies demonstrated that BMP, Wnt, and Nodal signaling are able to block neural differentiation of ESCs, mirroring findings from studies of zebrafish and Xenopus development (Finley et al., 1999; Wiles and Johansson, 1999; Kimelman and Griffin, 2000; Ikeda et al., 2005; Watanabe et al., 2005). That neurectoderm formation does indeed represent a default differentiation pathway for ESCs has since been confirmed using serum-free lowdensity embryoid body (EB) and monolayer culture differentiation systems where cell-cell interactions are minimized (Tropepe et al., 2001; Ying et al., 2003; Smukler et al., 2006). Although it is unclear what factors in serum are responsible for blocking neural differentiation (and inducing mesendoderm differentiation), the fact that the neural default can be restored by BMP, Wnt, or Nodal antagonists suggests that serum inhibits neural development by inducing secretion of proteins that activate these signaling pathways (Aubert et al., 2002; Gratsch and O’Shea, 2002; Pera et al., 2004; Watanabe et al., 2005). Addition of retinoic acid (RA) during the early phases of ESC differentiation can also promote neural differentiation (Meyer et al., 2004; Okada et al., 2004; Chiba et al., 2005). Again, al-

though the molecular mechanism by which RA exerts this affect has not been determined, it is possible that RA’s ability to induce expression of the Wnt antagonist, dkk1, may be important for RA-dependent neural differentiation (Verani et al., 2006). Finally, it is worth noting that once neurectodermal fate has been established, the same pathways whose signaling were blocked in order to achieve neural induction in the first instance are used to pattern the nascent neurectodermal to specific cell fates (Okada et al., 2004; Chiba et al., 2005; Irioka et al., 2005; Kawaguchi et al., 2005; Watanabe et al., 2005; Su et al., 2006). The use of the same signaling pathways to induce and then subsequently pattern cells of the nascent germ layers is a theme common to both neurectoderm and mesendoderm formation.

Mesendoderm induction Gene targeting studies have demonstrated that the secreted factors BMP4 (Winnier et al., 1995), Nodal (Conlon et al., 1994), Wnt3 (Liu et al., 1999), and FGF8 (Sun et al., 1999b) are required for primitive streak formation in the mouse. Studies using chimeric embryos, in which the expression of these genes is absent from either the embryonic or extraembryonic region, point to distinct roles for these proteins during germ layer formation and patterning. Analysis of chimeras of BMP4 null embryos and wild-type ESCs showed that BMP4 expression within extraembryonic ectoderm is required for the initiation of gastrulation (Fujiwara et al., 2001). Conversely, Nodal expression within the epiblast is required for gastrulation while its expression in extraembryonic endoderm regulates induction and patterning of anterior neural tissues (Varlet et al., 1997). Recent work suggests that BMP4, Nodal, and Wnt3 form a regulatory loop whereby Nodal signals from the epiblast maintain expression of BMP4 in the extraembryonic ectoderm, which in turn induces expression of Wnt3 in proximal epiblast (Ben-Haim et al., 2006). Intriguingly, these studies also showed that Nodal is unable to directly induce expression of either Brachyury or Wnt3, implying Nodal acts indirectly through up-regulation of extraembryonic BMP4, which in turn activates embryonic Wnt3 expression. Wnt3 maintains Nodal expression in the epiblast thus resulting in the establishment of a positive reinforcing feedback loop (Ben-Haim et al., 2006). Paralleling findings from studies in the embryo, experiments using ESCs differentiated in vitro suggest that BMP, Activin, and Wnt signals are all capable of initiating the

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formation of mesendoderm (Johansson and Wiles, 1995; Kubo et al., 2004; Park et al., 2004; D’Amour et al., 2005; Ng et al., 2005; Tada et al., 2005; Yasunaga et al., 2005; Gadue et al., 2006; Lindsley et al., 2006). By and large, findings from ESC differentiation studies can be rationalized in the context of developmental congruence between the in vitro and in vivo systems. For example, a number of reports have now shown that sustained Activin signaling promotes formation of definitive endoderm (DE) from ESCs (Kubo et al., 2004; D’Amour et al., 2005; Tada et al., 2005; Yasunaga et al., 2005; Gadue et al., 2006), consistent with the requirement for Nodal signaling in gut endoderm formation and with lineage tracing experiments showing that DE arises from the anterior primitive streak, a region of robust Nodal expression (Lowe et al., 2001; Lawson and Schoenwolf, 2003; Tam et al., 2003). Similarly, BMP4 promotes the formation of hematopoietic mesoderm (Johansson and Wiles, 1995; Wiles and Johansson, 1997; Li et al., 2001; Park et al., 2004; Ng et al., 2005) and cardiac mesoderm (Honda et al., 2006; Hosseinkhani et al., 2007), the latter in a concentration-dependent manner, reflecting findings from studies in Xenopus and zebrafish documenting the role of BMP4 as a morphogen (Dosch et al., 1997; Neave et al., 1997). Finally, it has recently been established that Nodal and Wnt signaling are both required for induction of primitive streak genes during ESC differentiation (Gadue et al., 2006), providing an in vitro correlate of genetic ablation studies in the developing mouse embryo (Conlon et al., 1994; Liu et al., 1999).

CONCLUSIONS

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Although the similarities between ESC differentiation and early mammalian development are compelling, one must also bear in mind that there is no a priori reason why cells must behave according to embryological principles. Indeed, the ability to maintain immortal, pluripotent ESCs is a poignant reminder that in vitro cultures can lead to the generation of nonphysiological cell types. Moreover, the outcome desired by the embryo is not the same as that sought by the experimenter. Whereas the aim of embryogenesis is to produce an animal, the aim of directed differentiation protocols is often to generate pure populations of a single cell type. Thus it is possible that many of the considerations that constrain the course of cell differentiation within the embryo may not necessarily apply to ESC differentiation. Nevertheless, because the em-

bryo has already mapped out differentiation pathways for the generation of every known cell type, it would seem prudent to begin with nature’s road map. In this regard, the most detailed maps are those generated from the study of frogs and fish, organisms that continue to provide new insights into the molecular mechanisms underlying vertebrate development.

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Germ Layer Induction in ESC

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Current Protocols in Stem Cell Biology

Formation and Hematopoietic Differentiation of Human Embryoid Bodies by Suspension and Hanging Drop Cultures

UNIT 1D.2

Chantal Cerdan,1 Seok Ho Hong,1 and Mickie Bhatia1 1

McMaster University, Hamilton, Ontario, Canada

ABSTRACT The in vitro aggregation of human embryonic stem cells (hESCs) into clusters termed embryoid bodies (EBs) allows for the spontaneous differentiation of cells representing endoderm, mesoderm, and ectoderm lineages. This stochastic process results however, in the generation of low numbers of differentiated cells, and can be enhanced to some extent by the addition of exogenous growth factors or overexpression of regulatory genes. In the authors’ laboratory, the use of hematopoietic cytokines in combination with the mesoderm inducer bone morphogenetic protein-4 (BMP-4) was able to generate up to 90% of CD45+ hematopoietic cells with colony-forming unit (CFU) activity. This unit describes two protocols that have been successfully applied in the authors’ laboratory for the generation of EBs in (1) suspension and (2) hanging drop (HD) cultures from enzymatically digested clumps of undifferentiated hESC colonies. Curr. Protoc. Stem C 2007 by John Wiley & Sons, Inc. Cell Biol. 3:1D.2.1-1D.2.16.
Keywords: human embryonic stem cells r embryoid body r suspension cultures r hanging drop cultures r differentiation r hematopoiesis

INTRODUCTION When withdrawn from their mouse embryonic fibroblast (MEF) feeder layers or FGFcontaining MEF–conditioned medium (MEFCM), which sustain their pluripotent state, human embryonic stem cells (hESCs) form spherical structures in suspension termed embryoid bodies (EBs) that comprise differentiated cells from the three germ layers (Chadwick et al., 2003, Cerdan et al., 2004; Tian et al., 2004; Wang et al., 2004; Zhan et al., 2004; Kim et al., 2005; Ng et al., 2005; Wang, 2005a,c; Zambidis et al., 2005; Bowles et al., 2006; Cameron et al., 2006). Alternatively, hESCs can be replated on various stromal cell layers (Kaufman et al., 2001; Tian et al., 2004; Narayan et al., 2005; Qiu et al., 2005; Vodyanik et al., 2005; Woll et al., 2005; Slukvin et al., 2006; Tian et al., 2006) known to support lineage development from somatic-derived populations. Somatic-derived populations refer to adult-derived populations. In the case of hematopoietic lineages, adult sources may be peripheral blood, umbilical cord blood, or bone marrow. This does not refer to the stromal cells. Both differentiation methodologies are largely adapted from methodologies used for mouse ESC (mESC) differentiation; they demonstrate similar efficiency in promoting hematopoietic development from hESCs; and they recapitulate temporal and spatial features of human embryonic development (Tavian et al., 1999, 2001; Oberlin et al., 2002). The spontaneous conversion of undifferentiated hESCs to blood lineages is generally a low-efficiency process and results in heterogeneous cell populations. Much of the literature on improving these efficiencies refers to the activation of specific developmentally relevant signaling pathways during EB development. Although more growth factors are required to support hematopoiesis in the Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1D.2.1-1D.2.16 Published online October 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01d02s3 C 2007 John Wiley & Sons, Inc. Copyright


1D.2.1 Supplement 3

Figure 1D.2.1 (A) and (B) show typical morphologies of EBs developing in suspension and hanging drop cultures, respectively. In both cultures, cystic EBs can be observed after day 7. Scale bar = 200 µm.

EB system compared to stroma cell–based differentiation of hESCs, both differentiation systems remain more effective in the presence of serum. However, serum may contain inhibitory factors, and differentiation efficiency has been described as being dependent on serum batch. There are no reports to date describing complete conversion of hESCs into hematopoietic lineages, and genetic selection methods as reported for mESCs are presently under development. A two-step process combining both the EB- and stromabased methodologies could maximize the efficiency of hematopoietic differentiation from hESCs, as recently suggested (Kim et al., 2005; Wang et al., 2005a). Altogether, these characteristics suggest that hESCs provide a unique resource for understanding the cellular and molecular basis of early human hematopoietic development or identifying genes that may cause abnormal development, in addition to possibly providing an unlimited supply of different lineages for replacement. This unit describes two methodologies, e.g., (1) suspension and (2) hanging drop cultures for the generation and hematopoietic differentiation from human EBs (see Fig. 1D.2.1). Both methodologies achieve EB generation from enzymatic digestion of hESC colonies resulting in the production of cellular clumps of variable sizes and compositions associated with cell death, which is likely enhanced by cavitation and apoptosis during EB development in culture (Coucouvanis and Martin, 1995). Unlike the mouse system, no current protocol for EB formation using either method has been optimized from single hESCs. While the protocols described here may be further optimized and modified depending on the study intended, they will likely remain important methods for differentiation of hESCs. An indication of which methods have been tried on which hESC lines with which relative efficiency is also provided. NOTE: All protocol steps should be performed at room temperature under sterile conditions in a Class II biological safety cabinet, unless otherwise mentioned. All incubations are performed in a humidified 37◦ C, 5% CO2 incubator, unless otherwise specified. Avoid repeated freeze-thaw cycles of all media/reagents/solutions stored at −80◦ C and −30◦ C. Formation and Hematopoietic Differentiation of Human Embryoid Bodies

NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic techniques should be used accordingly.

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Current Protocols in Stem Cell Biology

FORMATION AND HEMATOPOIETIC DIFFERENTIATION OF HUMAN EMBRYOID BODIES USING SUSPENSION CULTURES

BASIC PROTOCOL

Generally, most researchers have used suspension or methylcellulose cultures in order to differentiate adult or embryonic stem cells. In both suspension and hanging drop (HD) cultures, undifferentiated hESCs are allowed to grow in suspension where they aggregate to form three-dimensional structures termed embryoid bodies (EBs). During EB development, spontaneous differentiation toward hematopoietic lineages occurs in a very inefficient and uncontrolled way. Such differentiation can be enhanced by supplementing the EB medium with a cocktail of five cytokines and the mesoderm inducer BMP-4 for the entire period of culture (usually 15 to 22 days), as depicted in Table 1D.2.1. Medium and supplements are changed every 4 to 5 days and cells differentiate toward erythroid/myeloid phenotypes, as described elsewhere (Chadwick et al., 2003). Under such conditions, 10% to 20% of cells routinely become positive for the mature hematopoietic marker CD45 around day 15 of EB development. Therefore, this unit will focus on the description of the generation of EBs from clumps of hESCs using suspension and HD culture methods (see Alternate Protocol). In the authors’ hands, both protocols have been successfully used with H1, H9 (WiCell, http://www.wicell.org) cell lines propagated without antibiotics, using a Matrigel matrix, and MEFCM supplemented with basic fibroblast growth factor (bFGF, 8 ng/ml).

Materials hESC, undifferentiated clumps Matrigel (10-ml bottle; BD Biosciences, no. 353234; see recipe) Knockout DMEM (KO-DMEM; Invitrogen, no. 10829-018) Undifferentiated hESCs MEFCM (see recipe) Basic fibroblast growth factor (bFGF), recombinant human (see recipe) Collagenase IV (see recipe) EB medium (see recipe) Growth factors/cytokines: Stem cell factor (SCF), recombinant human (see recipe) Flt-3 ligand (Flt-3L), recombinant human (see recipe) Interleukin-3 (IL-3), recombinant human (see recipe) Interleukin-6 (IL-6), recombinant human (see recipe) Granulocyte colony–stimulating factor (G-CSF), recombinant human (300 µl bottle, Amgen, no. 3105100); aliquot 50 µl/tube and use it at 300 ng/µl (store no longer than 3 months at −30◦ C; store thawed aliquots no longer than 1 month at 4◦ C) Bone morphogenetic protein-4 (BMP-4), recombinant human (see recipe) Collagenase B (see recipe) Cell dissociation buffer, enzyme-free PBS-based (Invitrogen, no.13151-014) Iscove’s modified Dulbecco’s medium (IMDM) FACS buffer 100% fetal bovine serum (FBS) 6-well plates, tissue culture treated, flat bottom (VWR, cat. no. CA62406-161) 10-ml pipet 6-well ultra-low attachment plates, flat bottom (Fisher, cat. no. CS003471) 15-ml polypropylene tube 37◦ C water bath 200- or 1000-µl pipettor 40-µm cell strainer, nylon (BD Biosciences, no. 352340) Additional reagents and equipment for counting cells by trypan blue exclusion using a hemacytometer (UNIT 1C.3) Current Protocols in Stem Cell Biology

Embryonic and Extraembryonic Stem Cells

1D.2.3 Supplement 3

Prepare confluent and densely packed hESC colonies 1. At a time point ∼1 week before forming EBs, coat each well of a 6-well tissue culture–treated plate, with 0.75 to 1 ml of Matrigel at 1:6 dilution. 2. Aspirate the excess Matrigel, rinse each well of the plate with 2 ml KO-DMEM, aspirate, and seed undifferentiated hESCs (clumps of minimum 50 to 500 cells) in the 1:6 Matrigel precoated wells. 3. Feed cells daily with 3 ml MEFCM, prewarmed to 37◦ C, supplemented with fresh bFGF at 8 ng/ml to each well of 6-well plate. 4. At day 2, after addition of MEFCM + bFGF to the hESC culture, add 0.5 ml of 1:6 diluted Matrigel to each well, and gently swirl the plate to mix media. A 1:6 dilution of Matrigel is added to existing cultures of hESCs (at day 2 after passage). The exact mechanism of this effect is unknown—thickness of the hESC colonies is increased and the result on the hematopoietic output is better compared to no addition of Matrigel.

5. Incubate and proceed with the culture until 80% to 90% confluence is reached, with densely packed colonies. IMPORTANT NOTE: This may require longer than 7 days but <10 days. Cell density and thickness of the hESC colonies are critical for both maintaining the undifferentiated properties and achieving good EB formation. If >10 days are required, pass the cells at a 1:1 or even 2:1 (=“Back-passing”) ratios, onto fresh 1:6 Matrigel at day 10, and repeat, if necessary, until the above conditions are reached prior to proceeding with EB formation. Proper conditions for EB formation (cell density/thickness) can also be achieved by passaging the cells one week before on 1:15 dilution of Matrigel with no extra addition of Matrigel after passaging.

Form EBs 6. When proper density and thickness of the undifferentiated hESC colonies are reached (around day 7), aspirate the medium and add 0.5 ml of prewarmed collagenase IV (200 IU/ml) to each well. 7. Incubate for 3 to 15 min until colony edges slightly pull away from the well. Determine the appropriate incubation time by examining the colonies under the microscope. 8. Aspirate the collagenase IV. 9. Wash the residual collagenase IV with 2 ml of prewarmed KO-DMEM or EB medium per well and aspirate. 10. Add 2 ml of EB medium to each well. 11. Using a 10-ml pipet, gently scrape the bottom of each well in long “strips.” 12. Triturate VERY gently (ten times) using a 10-ml pipet. 13. Transfer the content of 2 wells of undifferentiated hESCs to 1 well of a 6-well ultra-low-attachment plate (ratio 2:1). This ratio can be 1.5:1. Total volume of EB medium per well is 4 ml.

14. Incubate overnight up to 24 hr to allow for EB formation. Formation and Hematopoietic Differentiation of Human Embryoid Bodies

This step is considered day 0 of EB differentiation.

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Promote hematopoietic development of EBs by hematopoietic cytokines and BMP-4 15. Transfer each well of EBs to an individual 15-ml polypropylene tube overnight to 24 hr post-EB formation. 16. Centrifuge 1 min at 129 × g, room temperature, or let settle a few minutes by gravity to separate EBs from debris and single cells. 17. Aspirate supernatant. Add 4 ml of EB medium supplemented with SCF, Flt3L, IL-3, IL-6, G-CSF, and BMP-4, or SCF, Flt3L, and BMP-4, to each tube. For specific quantities (stock and final concentrations, volumes) of cytokines and BMP-4, refer to Table 1D.2.1. No statistical difference between these two treatments is seen in the authors’ hands. This step is considered as day 1 of EB differentiation. EB differentiation can be maintained up to one month, with regular change of EB medium and cytokines.

18. Transfer each tube of EBs to the previous wells and incubate. 19. Change EB medium containing hematopoietic cytokines and BMP-4 every 4 to 5 days or earlier if medium changes color (orange/yellow). For a total of 15 days of differentiation, cultures are fed on days 1, 5, and 10.

Dissociate EBs 20. Transfer each well of EBs to an individual 15-ml polypropylene tube. These steps can be used for EB dissociation into single cells at any time of the differentiation period. The authors typically dissociate the EBs at two different time points: day 10 and day 15. Dissociation and analysis can be flexible around the two time points. Day 10 corresponds to the emergence of a precursor population called CD45− PFV (expressing PECAM-1, Flk-1, and VE-cadherin but devoid of CD45 expression). This population gives rise to CD45+ cells with a frequency of 10% to 20% around day 15 (Wang et al., 2004).

21. Centrifuge 3 min at 129 × g, room temperature. Aspirate supernatant. 22. Add 4 ml of collagenase B (0.4 IU/ml). Transfer to the previous well. Let EBs dissociate for 2 hr in a 37◦ C incubator. 23. Transfer the cells to a 15-ml polypropylene tube. Centrifuge for 10 min at 453 × g, room temperature. 24. Aspirate the collagenase B supernatant. 25. Add 2 ml of cell dissociation buffer and incubate for 10 min in a 37◦ C water bath. Table 1D.2.1 Supplementation of EB Medium with Cytokines and BMP-4 to Promote Hematopoietic Differentiation of EBs in Suspension or Hanging Drop Cultures

Growth factors/cytokines

Stock concentration

Final concentration

Volume for 4 ml of EB medium

BMP-4

50 ng/µl

25 ng/ml

2 µl

Flt-3L

250 ng/µl

300 ng/ml

4.8 µl

G-CSF

300 ng/µl

50 ng/ml

0.7 µl

IL-3

25 ng/µl

10 ng/ml

1.6 µl

IL-6

50 ng/µl

10 ng/ml

0.8 µl

SCF

1.5 × 10 ng/µl

300 ng/ml

0.8 µl

3

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26. Centrifuge for 10 min at 453 × g, room temperature. Aspirate the supernatant and depending on the number of EBs, resuspend in 100 to 500 µl of: a. IMDM for CFU plating or in vivo experiments. Add 100 µl IMDM for 25,000 to 100,000 EB cells. b. FACS buffer for flow cytometry analysis. Add 200 µl of FACS buffer for 10,000 to 100,000 EB cells. c. 100% FBS for cell sorting at a concentration of 10 ×106 EB cells minimum. 27. To achieve a single-cell suspension, gently triturate (50 to 70 times) with a 200-µl pipettor (set at 100 µl) or a 1000-µl pipettor (set at 200 to 450 µl) depending on the number of cells. 28. Filter through a sterile 40-µm cell strainer. 29. Count live and dead cells by trypan blue exclusion using a hemacytometer (UNIT 1C.3) and resuspend in the appropriate medium at the proper concentration, depending on the type of analysis (refer to step 26 for either CFU plating, FACS analysis, or in vivo transplantation). ALTERNATE PROTOCOL

FORMATION AND HEMATOPOIETIC DIFFERENTIATION OF HUMAN EMBRYOID BODIES USING HANGING DROP CULTURES Besides the suspension and methylcellulose methods, the hanging drop (HD) culture method can be selected for differentiation of stem cells and provides some unique conditions different from suspension and methylcellulose cultures. The most unique advantage of HD culture is to augment signaling between cells by increasing the cellular proximity within the small space of hanging droplet. Actually, there are some reports showing that HD culture is very useful to improve the rate of proliferation and differentiation of EBs generated using single cells of mESCs (Yamada et al., 2002; Zhao et al., 2005; Shang et al., 2006). Likewise, EBs can be generally formed from single cells and clumps of undifferentiated colonies of hESCs by suspension, but there is no optimal protocol for EB formation from single cells by HD culture. After ESC colonies are dissociated to single cells, they are not easily reaggregated without force such as spin down (Ng et al., 2005).

Materials hESC colonies in 6-well plates (see Basic Protocol) EB medium (see recipe) Collagenase IV (see recipe) Knockout DMEM (KO-DMEM; Invitrogen, cat. no.10829-018) 37◦ C water bath 5- or 10-ml serological pipet 60 × 15–mm polystyrene petri dish (Falcon, cat. no. 25382) 100 × 15–mm polystyrene petri dish (VWR, cat. no. 25384-088) 5- and 9-in. sterilized glass pipets (VWR, cat. no. 53283-914 and 14672-200) Rubber bulb (VWR, cat. no. 82024-562) 15-ml conical tube 6- and 24-well plates, ultra-low-attachment, flat bottom (Fisher, cat. no. CS003471 and CS003473)

Formation and Hematopoietic Differentiation of Human Embryoid Bodies

Prepare confluent and densely packed hESC colonies 1. Follow steps 1 to 5 of the Basic Protocol to prepare hESC colonies. Form EB in hanging droplet 2. Prewarm EB medium and collagenase IV in a 37◦ C water bath.

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Current Protocols in Stem Cell Biology

3. Aspirate hESC medium from a well with confluent and healthy hESC colonies. 4. Add 1 ml prewarmed collagenase IV and incubate for 3 to 15 min, until colony edges slightly pull away from the well. Determine the appropriate incubation time by examining the colonies under the microscope. 5. Aspirate collagenase IV and wash with 2 ml prewarmed KO-DMEM or EB medium per well. 6. Aspirate and add 2 ml of prewarmed EB medium. 7. Gently scrape the bottom of each well in long strips using a 5- or 10-ml serological pipet. 8. Triturate VERY gently (ten times) using a 5- or 10-ml serological pipet. 9. Add 5 to 6 ml (or 8 to 10 ml for a 100 ×15–mm polystyrene petri dish) of KO-DMEM or EB medium in a 60 × 15–mm petri dish for humidity. 10. Using a sterilized 9-in. glass pipet with rubber bulb, aspirate one clump of hESCs with EB medium. Large clumps contain ∼2000 to 3000 cells and small clumps contain between 300 and 500 cells. The volume of EB medium is variable depending on the size of clump, and generally 20 to 40 µl of medium is proper to hang the clumps.

11. Place drop with clump on the lid of a 60- or 100-mm petri dish (Fig. 1D.2.2). The optimal number is nine (3 × 3) droplets for 60-mm and twenty-five (5 × 5) droplets for 100-mm petri dishes.

12. Invert the lid with drops in one fluid motion over the bottom of the medium dish and incubate for 1 to 4 days according to the purpose of experiment. For instance, one day is sufficient to form EBs for hematopoietic differentiation in the authors’ laboratory.

Harvest EBs from hanging droplets 13. Prewarm EB medium in 37◦ C water bath prior to harvest EBs. 14. Take the dish from the incubator and transfer to the biological safety cabinet without shaking.

Figure 1D.2.2 (A) Shows a 9-in. sterilized glass pipet with rubber bulb to collect the clumps of hESCs and hanging droplets. (B) and (C) represent typical distributions of hanging droplets on the lid of 60- and 100-mm petri dishes, respectively. (D) Prior to inverting the lid to place it on the bottom portion of the dish, enough volume of KO-DMEM or EB medium (5 to 6 ml for a 60-mm dish and 8 to 10 ml for a 100-mm dish) must be added to the petri dish for maintenance of humidity. Embryonic and Extraembryonic Stem Cells

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15. Aspirate 1 to 2 ml of EB medium into a 5-in. glass pipet using a rubber bulb. 16. Invert the lid of the dish and drop 1 to 2 ml of EB medium onto the lid. 17. Collect EBs on the lid and transfer to a 15-ml conical tube. 18. Let settle by gravity until no EB clumps are found in supernatant. 19. Aspirate the supernatant and resuspend EBs in EB medium supplemented with appropriate factors depending on the specific cell lineages for differentiation (refer to Table 1D.2.1 for hematopoietic differentiation). 20. Transfer EBs to a 6- or 24-well ultra-low-attachment plate. Generally, EBs harvested from four and two 60-mm dishes are appropriate to transfer to one well of 6- and 24-well plates, respectively.

21. Continue incubation as long as required for optimal differentiation. EBs are usually grown in EB medium supplemented with hematopoietic cytokines and BMP-4 during 15 days. The medium should be replaced with fresh EB medium plus cytokines and BMP-4 every 3 to 5 days.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Collagenase B To prepare stock solution: Calculate the total amount of international units (IU) in each individual collagenase B bottle (100-mg bottle; Roche Diagnostics GmbH, cat. no. 1088-807) Dilute with KO-DMEM to make 4.0 IU per ml Filter through a 0.22-µm membrane Aliquot 1-ml stock solution into 15-ml polypropylene tubes Store at −30◦ C To prepare working solution (0.4 IU/ml), combine: 9 ml of KO-DMEM 1 ml of stock collagenase B solution Store up to 1 year at −30◦ C Thawed working solution of collagenase B can be stored at 4◦ C up to 1 week.

Collagenase IV

Formation and Hematopoietic Differentiation of Human Embryoid Bodies

To prepare stock solution: Calculate the total amount of IU in each individual collagenase IV bottle (1-g bottle; Invitrogen, cat. no. 17104-019) Dilute with KO-DMEM to make 10,000 IU per ml Aliquot 2 ml stock solution Store at −30◦ C To prepare working solution, combine: 98 ml of KO-DMEM 2 ml of stock collagenase IV solution Filter through a 0.22-µm membrane Store up to 1 year at −30◦ C Thawed working solution of collagenase IV can be stored at 4◦ C up to 1 week.

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Current Protocols in Stem Cell Biology

EB medium For 100 ml, combine: 20 ml non-heat-inactivated fetal bovine serum, FBS (20% v/v final, Hyclone, cat. no. 30071-03) 0.5 ml 200 mM L-glutamine (1 mM final, Invitrogen, cat. no. 25030-081) 80 ml KO-DMEM (Invitrogen, cat. no. 10829-018) 7 µl 1.43 M 2-mercaptoethanol (0.1 mM final, Sigma, cat. no. M7522) 1 ml 10 mM non-essential amino acids (1% v/v final, Invitrogen, cat. no. 1140-050) Filter through a 0.22-µm sterile cellulose acetate membrane (150-ml and 500-ml; Fisher, cat. no. 09-761-119 and 09-761-5) Store no longer than 14 days at 4◦ C Once thawed, 2-mercaptoethanol can be stored up to 1 month at 4◦ C and L-glutamine can be stored up to 2 to 4 weeks at 4◦ C. Non-essential amino acids are stable at 4◦ C up to the date of expiration.

Growth factors/cytokines bFGF To prepare working solution (10 µg/ml): Dissolve each vial of bFGF (10-µg vial; Invitrogen, cat. no. 13256-029) in 1 ml D-PBS (Invitrogen, cat. no. 14190-144) containing 0.1% (w/v) BSA (add 30 µl of 30% w/v BSA into 10 ml D-PBS). Prefilter Ultra free-MC 0.22-µm filter (Millipore, cat. no. UFC30GVOS) with 3 ml of D-PBS containing 10% (w/v) BSA (combine 1 ml of 30% w/v BSA with 2 ml D-PBS). Filter bFGF working solution. Divide filtered bFGF into 200-µl aliquots using sterile Eppendorf tubes. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. BMP-4 To prepare stock solution (1000 µg/ml): Add 300 µl of D-PBS containing 2% (v/v) heat-inactivated FBS to the bottle of BMP-4 (300-µg bottle; R&D, cat. no. 314-BP), pipet, and aliquot 50 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. To prepare working solution (50 µg/ml): Add 950 µl of D-PBS containing 2% (v/v) heat-inactivated FBS to 50 µl of stock solution, pipet, and aliquot 50 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. Flt-3L To prepare working solution (250 µg/ml): Add 1 ml of D-PBS containing 2% (v/v) heat-inactivated FBS to the bottle of Flt-3L (250-µg bottle; R&D, cat. no. 308FK/CF), pipet, and aliquot 100 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. G-CSF Aliquot 50 µl/tube, and use as it is at 300 ng/µl. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. IL-3 To prepare working solution (25 µg/ml): Add 2 ml of D-PBS containing 2% (v/v) heat-inactivated FBS to the bottle of IL-3 (50-µg bottle; R&D, cat. no. 203-IL), pipet, and aliquot 100 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C.

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IL-6 To prepare working solution (50 µg/ml): Add 1 ml of D-PBS containing 2% (v/v) heat-inactivated FBS to the bottle of IL-6 (50-µg bottle; R&D, cat. no. 206-IL), pipet, and aliquot 50 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C. SCF (stem cell factor) To prepare working solution (1500 µg/ml): Add 660 µl of D-PBS containing 2% (v/v) heat-inactivated FBS to the bottle of SCF (1000-µg bottle; R&D, cat. no. 255SC/CF), pipet, and aliquot 50 µl/tube. Store no longer than 3 months at −30◦ C. Store thawed aliquots no longer than 1 month at 4◦ C.

Matrigel To prepare stock solution: Thaw Matrigel slowly to avoid gel formation (overnight at 4◦ C). Add 10 ml of cold KO-DMEM (4◦ C). Put Matrigel bottle and 20 tubes (15-ml polypropylene conical tubes) on ice. Pipet 20 times to mix Matrigel and KO-DMEM medium. Make 1 ml/tube aliquots and store up to 3 months from date of shipment at −30◦ C. To prepare and coat wells with 1:15 working solution: Thaw one tube of stock Matrigel at 4◦ C overnight or within 1 to 2 hr into a container with cold water to avoid polymerization (which occurs just above 4◦ C). Add 5 ml of cold KO-DMEM and pipet 10 times. Add 9 ml of cold KO-DMEM and pipet 10 times. Add 0.75 to 1 ml of working solution to one well of a 6-well plate. Spread evenly without forming bubbles. Wrap the plate with plastic film. Incubate the plate overnight at 4◦ C or for 2 hr at room temperature to allow polymerization. Use coated plates immediately or store no longer than 7 days at 4◦ C. To prepare and coat wells with 1:6 dilution: For the passage of hESCs one week prior to EB formation, proceed the same way as for 1:15 working solution but add only 5 ml of cold KO-DMEM into 1 ml Matrigel aliquots. Once diluted, Matrigel can be stored at 4◦ C no longer than a week.

2-mercaptoethanol To prepare 1.43 M 2-mercaptoethanol, combine: 18 ml of D-PBS, Ca2+ and Mg2+ free (Invitrogen, cat. no. 14190-144) 2 -ml of 14.3 M 2-mercaptoethanol Store up to 3 months at −30◦ C in 0.25-ml aliquots Store thawed 1.43 M 2-mercaptoethanol up to 1 month at 4◦ C Mouse embryonic fibroblast conditioned medium (MEFCM) To prepare 500 ml, combine: 392.5 ml KO-DMEM (80% v/v final; see recipe) 2.5 ml L-glutamine (1 mM final) 35 µl 1.43 M 2-mercaptoethanol (0.1 mM final; see recipe) 5 ml non-essential amino acids (1% v/v final) 100 ml KO-SR (20% v/v final; Invitrogen, cat. no. 10828-028) Filter through a 0.22-µm sterile cellulose acetate membrane Store up to 14 days at 4◦ C Immediately prior to use, add 200 µl of bFGF (working solution, 10 ng/µl) Formation and Hematopoietic Differentiation of Human Embryoid Bodies

continued

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Feed irradiated (40 Gy) MEF cells and collect medium daily up to 10 days. Pool all collected MEFCM, filter through a 0.22-µm sterile cellulose acetate membrane (500 ml volume). Store aliquots up to 6 months at −80◦ C. Store thawed MEFCM no more than 7 days at 4◦ C. MEFCF quality is crucial for maintenance of undifferentiated hESCs and subsequent formation and hematopoietic development of EBs. Since MEFCM quality varies among different batches of MEF cells and MEFCM preparations, assessing each batch of MEFCM prior to use is recommended. Every new batch of MEFCM is compared to the batch currently used on the hESC cultures by assessing (weekly) the morphology, growth rate, and phenotype of the cultures during a minimum of 3 weeks. Phenotypic analysis includes detection of SSEA-4, SSEA-3, and Oct-4 antigens.

COMMENTARY Background Information Suspension EBs Human EB formation varies in several technical ways from its mouse counterpart. In contrast to mouse EBs that initiate from single cells, development of human EBs typically involves clusters of cells that acquire the best differentiated properties when treated with media including tested lots of FBS, and are more difficult to achieve from single cells. Each new lot of FBS is tested against the one currently in use for EB differentiation. This means that differentiation of hematopoietic cells from EBs cultured in media plus the new FBS lot is analyzed by flow cytometry at days 10 and 15 for the frequency and total numbers of the CD45− PFV and CD45+ populations, respectively. If the frequencies and total numbers of both populations are similar between the two lots of FBS (new and current), then the new lot of FBS is ordered in as large quantities as possible— otherwise, another lot is tested. At present, three main methods of EB formation in suspension cultures have been reported for the induction of hematopoietic differentiation from hESCs. These include static suspension cultures as described here, stirred suspension cultures (Cameron et al., 2006), and spin EBs (Ng et al., 2005). Although the three methods retain comparable potential to generate hematopoietic progenitors from hESCs in terms of both kinetics and frequencies, they provide inherent and unique advantages and weaknesses. Static suspension cultures have the propensity to generate EBs with heterogeneous morphology and size, leading to inefficient and uncontrolled cellular differentiation. Although the morphology has been shown to have little impact on hESC-derived hematopoietic differentiation (Cameron et al., 2006), the size appears more critical with large- or medium-sized EBs giving the best yields of hematopoietic progenitor cells (S. H. Hong, unpub. observ.). Stirred suspension cultures tend to generate Current Protocols in Stem Cell Biology

more uniform EBs, with respect to their diameter and morphology, and result in greater cellular expansion than static cultures. Therefore, they provide a foundation toward large-scale production of hESC-derived progenies. However, as a result of exposure to greater shear stress, increased production of cellular debris is also observed in this system (Cameron et al., 2006). Through the aggregation of known numbers of hESCs by centrifugation, the spin EB method promotes the synchronous development of EBs of uniform size, and provides the minimum requirements of hESC numbers for optimum differentiation (Ng et al., 2005). Based on single-cell dissociation, this method paves the way for improving the introduction of genetic modifications in hESC-derived progenies and monitoring directed differentiation. However, EBs formed in suspension cultures are more amenable to scalable cell production of differentiated cell types than those developing in hanging drops, although they are prone to more agglomeration after the initial stages of formation (Dang et al., 2004). Hanging drop EBs Since the hanging drop technique was developed by Harrison in 1907 (Harrison, 1907), it has been widely used during the last century and several types of HD techniques have been developed for distinct purposes. Originally, the HD method was used to observe living developing cells (Gliozzi, 1959; Andrew and Gabie, 1969). It consisted of placing a small droplet of medium and cells on a coverslip, then inverting the coverslip over a depression slide so that the bottom of the droplet did not contact the slide itself. The HD method was then modified to examine the behavior of cells by placing drops containing specific cell types on the lid of a petri dish (Niederman and Armstrong, 1972; Armstrong and Armstrong, 1978). Different plasticwares, such as 60- or 100-mm culture dishes and multi-well culture plates, (Terasaki plate and mini-tray) have

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1D.2.11 Supplement 3

been used with this method (Hopkins and Das, 1973; Noren et al., 2006). This method provides different conditions compared with routine cell cultures in which cells are grown attached to the bottom of a dish/plate. First, cells cultured in HD avoid the cellular distortion typically seen when cells are allowed to attach to the surface of a dish/plate. Second, signaling between cells is promoted by increasing the cellular proximity within the small space of hanging droplets (Gutierrez et al., 2005). There are some reports showing a positive effect of such conditions on lymphoid B- and T-cell development (Sagara et al., 1997; Tokoro et al., 1998) and preimplantation embryonic development (Potter and Morris, 1985; Spindler et al., 2000; Swain et al., 2001). Since the derivation of hESCs in 1998 (Thomson et al., 1998), only one study applied the HD methodology to the differentiation of cardiomyocytes from clumps of undifferentiated hESCs (Yoon et al., 2006). However, there is no optimal protocol for HD culture using single cells in hESCs unlike mESCs (Dang et al., 2002; Kurosawa et al., 2003).

Critical Parameters

Formation and Hematopoietic Differentiation of Human Embryoid Bodies

Suspension EBs The density of the undifferentiated hESC culture is critical as well as the thickness of the individual colonies. This is improved by passaging the undifferentiated cells onto 1:6 dilution of Matrigel 1 week prior to EB formation, followed by the addition of 0.5 ml of 1:6 diluted Matrigel per well of a 6-well plate to the feeding medium at day 2 or 3 after passaging. Another important factor is the batch of non-heat-inactivated FBS used to prepare the EB medium. It is recommended that one test different batches for EB differentiation efficiency and choose the best performer. Similarly, the authors recommend performing quality control testing on every batch of MEFCM produced. The authors usually assess the expression of undifferentiated markers (such as SSEA-3, Oct3/4) by flow cytometry after a 3-week culture period of the hESCs with both the current and new batch of MEFCM. The authors noticed (unpub. observ.) that undifferentiated hESCs propagated in MEFCM (supplemented with 8 ng/ml bFGF) gave rise to greater and more reproducible hematopoietic differentiation from EB suspension cultures than cells cultured in SR (supplemented with 40 ng/ml bFGF; Wang et al., 2005b) or XVivo-10 (Li et al., 2005; supplemented with 80 to 100 ng/ml bFGF) media. In

addition, hESCs cultured in XVivo-10 medium maintained the ability to differentiate toward hematopoietic lineages for only 10 to 12 weeks (C. Cerdan, unpub. observ.). With regard to the efficiency of hematopoietic differentiation of EBs during suspension cultures but not their formation, a significant variability between hESC lines (in our hands, H9 is usually superior to H1), experiments, and experimenters was observed. Hanging drop EBs The most critical factors for successful HD culture using clumps of undifferentiated hESC colonies are the choice of a proper dish/plate, diameter of pipet to collect the clumps, and volume and number of droplets placed on the lid. The best results can be achieved by placing 3 × 3 droplets of 20 to 30 µl on the lid of a 60 × 15–mm petri dish and 5 × 5 droplets of 20 to 30 µl on the lid of a 100 × 15–mm petri dish. For the maintenance of humidity and structure of hanging droplets, each dish/plate must contain enough volume of KO-DMEM or EB medium in the bottom of the dish. Keeping the original shape of droplets during long-term culture periods is also critical for successful EB generation. Again, this can be achieved by choosing a dish/plate made of plastic which enables droplets to adopt the right shape and stay put [e.g., 60 × 15–mm polystyrene petri dish (cat. no. 25382) and 100 × 15–mm polystyrene petri dish (cat. no. 25384-088) supplied from VWR]. Since the enzymatic digestion and scraping of hESC colonies result in the generation of clumps of very different sizes, it is important to choose a pipet with the appropriate diameter. [Sterile aerosol filter pipet tips (VWR) of 1-ml volume can be used to load small clumps. A 5- or 9-in. glass pipet is adequate for loading large clumps.] To avoid agglomeration together of the droplets on the lid of the dish, both the volume and number of droplets placed on the lid must be tightly controlled.

Troubleshooting Suspension EBs See Table 1D.2.2 for a troubleshooting guide for EB formation by suspension culture. Hanging drop EBs See Table 1D.2.3 for a troubleshooting guide for EB formation by hanging drop culture.

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Current Protocols in Stem Cell Biology

Table 1D.2.2 Troubleshooting Guide for Formation of EBs in Suspension Cultures

Problem

Possible caus

Solution

Lack or weak hematopoietic differentiation

Insufficient density/thickness of the hESC colonies

Do not form EBs when density and thickness of the hESC colonies are not optimal: <80% -90% confluence or colonies too thin. Pre-seed the cells on 1:6 Matrigel prior to EB formation.

EB medium too old or insufficiently changed

Use EB medium for no longer than 14 days and monitor color changes (change before it gets yellow).

Maintenance of the undifferentiated hESC culture too long

Restart from a fresh thaw.

Absence or improper ratio of fibroblast-like cells to undifferentiated cells

Wait for the proper ratio and the good appearance of the hESC colonies or restart from a fresh thaw.

hESC colonies do not detach easily from the well after collagenase IV treatment or EBs are too small

Insufficient time of Monitor dissociation time under the microscope or use a incubation with collagenase different batch of collagenase IV. IV Colonies were not rolled in long strips

Make ribbons from one edge of the well to the opposite.

Pipetting too vigorous

Pipet very gently and only a few times.

Lot of cellular debris just after EB formation performed too Monitor the hESC colony density. EB formation late after passaging No washing after collagenase IV incubation

Wash once after collagenase dissociation.

Pipetting too vigorous

Pipet gently and only a few times during EB formation.

Table 1D.2.3 Troubleshooting Guide to the Formation of EBs in Hanging Drops

Problem

Cause

Hanging droplet is flattened Too large hanging droplet or use of improper petri dish/plate

Solution Reduce the volume of hanging droplet or select a dish/plate with proper plastic.

hESC colonies stick to the lid after hanging drop formation

Hanging droplets are too small or Put the proper volume and number of hanging too flat droplets or select the proper dish/plate.

Hanging droplets run when the lid is inverted

Use of serum-free medium or a fluid motion

Use medium supplemented with 10%-20% serum or invert the lid gently in one fluid motion.

Hanging droplets adhere to the lid

Too many droplets on the lid

Reduce the number of hanging droplets. Recommend 3 × 3 droplets for a 60 × 15-mm dish and 5 × 5 droplets for a 100 × 15-mm dish.

Hanging droplets Too narrow spacing between agglomerate together on the droplets or improper movement lid of droplets

Keep the proper distance between droplets or invert the lid in one fluid motion.

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Anticipated Results Suspension EBs In the setting of hematopoietic differentiation from EB suspension cultures, the cells remain typically uncommitted to the hematopoietic cell fate prior to day 10 of EB development, after which they progressively express the pan-leukocyte marker CD45. By day 15, ∼10% to 20% of all cells within EBs treated with cytokines that include SCF, Flt-3L, (IL-3, IL-6, G-CSF are optional) and BMP-4 express CD45. No statistical difference was observed between the use of SCF, Flt3L, and BMP-4 or SCF, Flt-3L, BMP-4, IL-3, IL-6, and G-CSF (C. Cerdan, unpub. observ.). Similarly to their somatic counterparts, differentiated hESCs with hematopoietic progenitor capacity revealed by CFU assay, are found to be restricted to the CD45+ population. Continuous treatment of the EBs up to day 22 further increases the frequency of the CD45+ population (up to 90%), without changing the CFU plating efficiency (1 in 260). Colonies arising from replating of EBs in methylcellulose standard assay demonstrate the characteristic morphology and distribution of CFU subtypes, including macrophage, granulocyte, erythrocyte, as well as bipotent and multipotent colonies. BMP-4 has little effect on hematopoietic progenitor formation, but enhances (3.5-fold) the self-renewal capacity of the progenitors. Interestingly, ∼10% to 20% of EBs at days 10 to 13 still contain Oct3/4+ cells (L. Li, unpub. observ.), indicating the lack of synchrony of this differentiation system.

Formation and Hematopoietic Differentiation of Human Embryoid Bodies

Hanging drop EBs Finding an optimal differentiation protocol using hESCs often requires the comparison of various protocols such as suspension versus methylcellulose or suspension versus hanging drop. Using clumps of hESC colonies, a sideby-side comparison of suspension and HD cultures for hematopoietic differentiation was performed in the authors’ laboratory. The HD method was successfully tested and showed a comparable efficiency of hematopoietic differentiation (both kinetic and frequencies of the different progenitor populations) as the suspension culture. Another advantage of the HD method from hESC clumps is that it allows partial control on the size and morphology of the EBs, which might be helpful for analyzing the EB potential in relation to distinct properties of undifferentiated hESC colonies. For the authors, using the HD method, large and medium size EBs appear to differentiate more efficiently toward hematopoietic lin-

eages than small EBs. The frequency of CD45+ and CD34+ /CD45+ cells in large and medium versus small-sized EBs (the authors calculated the diameter of the EBs by averaging the longest and shortest lengths) at day 15 of EB development are significantly different (higher in large and medium versus small EBs; S. H. Hong, unpub. observ.).

Time Considerations Suspension EBs The amount of time required for EB formation by suspension culture is comparatively shorter than by the HD method and can be divided into two phases. The first phase prepares the undifferentiated hESC culture to allow for the second phase of EB formation. The goal of the first phase is to achieve 80% to 90% confluent and densely packed hESC colonies, which usually requires 6 to 8 days depending on the proliferation rate and the ratio of undifferentiated hESCs to fibroblast-like cells which must be higher than 50/50. The fibroblast-like cells are derived from the hESC cultures. These cells do not express SSEA-3, unlike the cells forming the colonies. However, these cells do express Nanog and Oct-4 and retain pluripotent potential, as assessed by teratoma formation (Stewart et al., 2006; Bendall et al., 2007). It may take a little longer to achieve this goal but always <10 days. The second phase starts with collagenase IV digestion of the hESC colonies to break them down into smaller clumps and scrape them into long ribbons from the bottom of the well. The incubation time with collagenase IV may vary between 5 and 20 min (average of 10 min) depending on the extent of cellular detachment from the edge of the well and between the colonies, as well as the batch of enzyme. Thus, if two wells of a 6-well plate are used to form one 6-well of EBs, it typically requires <5 min from the aspiration of collagenase IV to the transfer of the hESC ribbons in the ultralow attachment well. The phase of hematopoietic differentiation usually takes between 15 and 22 days for completion but can be stopped at any time point for different experimental purposes. Hanging drop EBs The amount of time required for EB formation by HD is comparatively longer than suspension culture and is also divided into two phases. The duration of the first phase, which prepares confluent and densely packed hESC colonies prior to EB formation, has been described in the EB suspension culture protocol.

1D.2.14 Supplement 3

Current Protocols in Stem Cell Biology

For the second phase of EB formation from hESC clumps, the incubation time with collagenase IV has also been commented upon. The collection of one clump of hESCs and its transfer to the lid of a dish/plate usually takes 5 to 10 sec. Thus, if one well of a 6-well plate of undifferentiated hESCs is used for EB formation, ∼100 clumps can be placed on lids, with about 10 to 12 lids of 60-mm dishes or 4 to 5 lids of 100-mm dishes in ∼1 hr. The time for harvesting EBs from droplets is relatively short and requires 1 to 2 min per 60-mm dish or 2 to 3 min per 100-mm dish. For hematopoietic differentiation, it usually takes between 15 and 22 days to obtain >10% CD45+ cells and around 10 days to obtain the precursor population CD45negPFV (Wang et al., 2004).

Literature Cited Andrew, A. and Gabie, V. 1969. Hanging drop culture of Xenopus laevis neural crest: Acta Embryol. Exp. (Palermo) 2:123-136. Armstrong, M.T. and Armstrong, P.B. 1978. Cell motility in fibroblast aggregates: J. Cell Sci. 33:37-52.

scalable embryonic stem cell differentiation culture. Stem Cells 22:275-282. Gliozzi, M.A. 1959. Growth in culture in vitro in hanging drop alternatively transplanted & renourished. Boll. Soc. Ital. Biol. Sper. 35:377379. Gutierrez, L., Lindeboom, F., Ferreira, R., Drissen, R., Grosveld, F., Whyatt, D., and Philipsen, S. 2005. A hanging drop culture method to study terminal erythroid differentiation. Exp. Hematol. 33:1083-1091. Harrison, R.G. 1907. Observations on the living developing nerve fibres. Proc. Soc. Exp. Bio. Med. 4:140-143. Hopkins, R. and Das, P.C. 1973. A tanned cell haemagglutination test for the detection of hepatitis-associated-antigen (Au-Ag) and antibody (anti-Au). Br. J. Haematol. 25:619-629. Kaufman, D.S., Hanson, E.T., Lewis, R.L., Auerbach, R., and Thomson, J.A. 2001. Hematopoietic colony-forming cells derived from human embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 98:10716-10721. Kim, S.J., Kim, B.S., Ryu, S.W., Yoo, J.H., Oh, J.H., Song, C.H., Kim, S.H., Choi, D.S., Seo, J.H., Choi, C.W., Shin, S.W., Kim, Y.H., and Kim, J.S. 2005. Hematopoietic differentiation of embryoid bodies derived from the human embryonic stem cell line SNUhES3 in co-culture with human bone marrow stromal cells. Yonsei Med. J. 46:693-699.

Bendall, S.C., Stewart, M.H., Menendez, P., George, D., Vijayaragavan, K., WerbowetskiOgilvie, T., Ramos-Mejia, V., Rouleau, A., Yang, J., Bosse, M., Lajoie, G., and Bhatia, M. 2007. IGF and FGF cooperatively establish the regulatory stem cell niche of pluripotent human cells in vitro. Nature 448:1015-1021.

Kurosawa, H., Imamura, T., Koike, M., Sasaki, K., and Amano, Y. 2003. A simple method for forming embryoid body from mouse embryonic stem cells. J. Biosci. Bioeng. 96:409-411.

Bowles, K.M., Vallier, L., Smith, J.R., Alexander, M.R., and Pedersen, R.A. 2006. HOXB4 overexpression promotes hematopoietic development by human embryonic stem cells. Stem Cells 24:1359-1369.

Li, Y., Powell, S., Brunette, E., Lebkowski, J., and Mandalam, R. 2005. Expansion of human embryonic stem cells in defined serumfree medium devoid of animal-derived products. Biotechnol. Bioeng. 91:688-698.

Cameron, C.M., Hu, W.S., and Kaufman, D.S. 2006. Improved development of human embryonic stem cell-derived embryoid bodies by stirred vessel cultivation: Biotechnol. Bioeng. 94:938-948.

Narayan, A.D., Chase, J.L., Lewis, R.L., Tian, X., Kaufman, D.S., Thomson, J.A., and Zanjani, E.D. 2005. Human embryonic stem cell-derived hematopoietic cells are capable of engrafting primary as well as secondary fetal sheep recipients. Blood 107:2180-2183.

Cerdan, C., Rouleau, A., and Bhatia, M. 2004. VEGF-A165 augments erythropoietic development from human embryonic stem cells. Blood 103:2504-2512. Chadwick, K., Wang, L., Li, L., Menendez, P., Murdoch, B., Rouleau, A., and Bhatia, M. 2003. Cytokines and BMP-4 promote hematopoietic differentiation of human embryonic stem cells. Blood 102:906-915. Coucouvanis, E. and Martin, G.R. 1995. Signals for death and survival: a two-step mechanism for cavitation in the vertebrate embryo. Cell 83:279287. Dang, S.M., Kyba, M., Perlingeiro, R., Daley, G.Q., and Zandstra, P.W. 2002. Efficiency of embryoid body formation and hematopoietic development from embryonic stem cells in different culture systems: Biotechnol. Bioeng. 78:442-453. Dang, S.M., Gerecht-Nir, S., Chen, J., ItskovitzEldor, J., and Zandstra, P.W. 2004. Controlled, Current Protocols in Stem Cell Biology

Ng, E.S., Davis, R.P., Azzola, L., Stanley, E.G., and Elefanty, A.G. 2005. Forced aggregation of defined numbers of human embryonic stem cells into embryoid bodies fosters robust, reproducible hematopoietic differentiation. Blood 106:1601-1603. Niederman, R. and Armstrong, P.B. 1972. Is abnormal limb bud morphology in the mutant talpid 2 chick embryo a result of altered intercellular adhesion? Studies employing cell sorting and fragment fusion. J. Exp. Zool. 181:17-32. Noren, N.K., Foos, G., Hauser, C.A., and Pasquale, E.B. 2006. The EphB4 receptor suppresses breast cancer cell tumorigenicity through an Abl-Crk pathway. Nat. Cell Biol. 8:815-825. Oberlin, E., Tavian, M., Blazsek, I., and Peault, B. 2002. Blood-forming potential of vascular endothelium in the human embryo. Development 129:4147-4157.

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Potter, S.W. and Morris, J.E. 1985. Development of mouse embryos in hanging drop culture. Anat. Rec. 211:48-56.

1998. A mouse carrying genetic defect in the choice between T and B lymphocytes. J. Immunol. 161:4591-4598.

Qiu, C., Hanson, E., Olivier, E., Inada, M., Kaufman, D.S., Gupta, S., and Bouhassira, E.E. 2005. Differentiation of human embryonic stem cells into hematopoietic cells by coculture with human fetal liver cells recapitulates the globin switch that occurs early in development. Exp. Hematol. 33:1450-1458.

Vodyanik, M.A., Bork, J.A., Thomson, J.A., and Slukvin, I.I. 2005. Human embryonic stem cellderived CD34+ cells: Efficient production in the coculture with OP9 stromal cells and analysis of lymphohematopoietic potential. Blood 105:617626.

Sagara, S., Sugaya, K., Tokoro, Y., Tanaka, S., Takano, H., Kodama, H., Nakauchi, H., and Takahama, Y. 1997. B220 expression by T lymphoid progenitor cells in mouse fetal liver. J. Immunol. 158:666-676. Shang, L.L., Dudley, S.C. Jr., and Pfahnl, A.E. 2006. Analysis of arrhythmic potential of embryonic stem cell-derived cardiomyocytes. Methods Mol. Biol. 330:221-231. Slukvin, I.I., Vodyanik, M.A., Thomson, J.A., Gumenyuk, M.E., and Choi, K.D. 2006. Directed differentiation of human embryonic stem cells into functional dendritic cells through the myeloid pathway. J. Immunol. 176:2924-2932. Spindler, R.E., Pukazhenthi, B.S., and Wildt, D.E. 2000. Oocyte metabolism predicts the development of cat embryos to blastocyst in vitro. Mol. Reprod. Dev. 56:163-171. Stewart, M.H., Bosse, M., Chadwick, K., Menendez, P., Bendall, S.C., and Bhatia, M. 2006. Clonal isolation of hESCs reveals heterogeneity within the pluripotent stem cell compartment. Nat. Methods 3:807-815. Swain, J.E., Bormann, C.L., and Krisher, R.L. 2001. Development and viability of in vitro derived porcine blastocysts cultured in NCSU23 and G1.2/G2.2 sequential medium. Theriogenology 56:459-469. Tavian, M., Hallais, M.F., and Peault, B. 1999. Emergence of intraembryonic hematopoietic precursors in the pre-liver human embryo. Development 126:793-803. Tavian, M., Robin, C., Coulombel, L., and Peault, B. 2001. The human embryo, but not its yolk sac, generates lympho-myeloid stem cells: Mapping multipotent hematopoietic cell fate in intraembryonic mesoderm. Immunity 15:487-495. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Tian, X., Morris, J.K., Linehan, J.L., and Kaufman, D.S. 2004. Cytokine requirements differ for stroma and embryoid body-mediated hematopoiesis from human embryonic stem cells. Exp. Hematol. 32:1000-1009.

Formation and Hematopoietic Differentiation of Human Embryoid Bodies

Tian, X., Woll, P.S., Morris, J.K., Linehan, J.L., and Kaufman, D.S. 2006. Hematopoietic engraftment of human embryonic stem cell-derived cells is regulated by recipient innate immunity. Stem Cells 24:1370-1380. Tokoro, Y., Sugawara, T., Yaginuma, H., Nakauchi, H., Terhorst, C., Wang, B., and Takahama, Y.

Wang, L., Li, L., Shojaei, F., Levac, K., Cerdan, C., Menendez, P., Martin, T., Rouleau, A., and Bhatia, M. 2004. Endothelial and hematopoietic cell fate of human embryonic stem cells originates from primitive endothelium with hemangioblastic properties. Immunity 20:31-41. Wang, J., Zhao, H.P., Lin, G., Xie, C.Q., Nie, D.S., Wang, Q.R., and Lu, G.X. 2005a. In vitro hematopoietic differentiation of human embryonic stem cells induced by co-culture with human bone marrow stromal cells and low dose cytokines. Cell Biol. Int. 29:654-661. Wang, L., Li, L., Menendez, P., Cerdan, C., and Bhatia, M. 2005b. Human embryonic stem cells maintained in the absence of mouse embryonic fibroblasts or conditioned media are capable of hematopoietic development. Blood 105:45984603. Wang, L., Menendez, P., Shojaei, F., Li, L., Mazurier, F., Dick, J.E., Cerdan, C., Levac, K., and Bhatia, M. 2005c. Generation of hematopoietic repopulating cells from human embryonic stem cells independent of ectopic HOXB4 expression: J. Exp. Med. 201:1603-1614. Woll, P.S., Martin, C.H., Miller, J.S., and Kaufman, D.S. 2005. Human embryonic stem cell-derived NK cells acquire functional receptors and cytolytic activity. J. Immunol. 175:5095-5103. Yamada, T., Yoshikawa, M., Kanda, S., Kato, Y., Nakajima, Y., Ishizaka, S., and Tsunoda, Y. 2002. In vitro differentiation of embryonic stem cells into hepatocyte-like cells identified by cellular uptake of indocyanine green. Stem Cells 20:146-154. Yoon, B.S., Yoo, S.J., Lee, J.E., You, S., Lee, H.T., and Yoon, H.S. 2006. Enhanced differentiation of human embryonic stem cells into cardiomyocytes by combining hanging drop culture and 5-azacytidine treatment. Differentiation 74: 149-159. Zambidis, E.T., Peault, B., Park, T.S., Bunz, F., and Civin, C.I. 2005. Hematopoietic differentiation of human embryonic stem cells progresses through sequential hematoendothelial, primitive, and definitive stages resembling human yolk sac development. Blood 106:860-870. Zhan, X., Dravid, G., Ye, Z., Hammond, H., Shamblott, M., Gearhart, J., and Cheng, L. 2004. Functional antigen-presenting leucocytes derived from human embryonic stem cells in vitro. Lancet 364:163-171. Zhao, X., Teng, R., Asanuma, K., Okouchi, Y., Johkura, K., Ogiwara, N., and Sasaki, K. 2005. Differentiation of mouse embryonic stem cells into gonadotrope-like cells in vitro. J. Soc. Gynecol. Investig. 12:257-262.

1D.2.16 Supplement 3

Current Protocols in Stem Cell Biology

Directed Differentiation of Human Embryonic Stem Cells as Spin Embryoid Bodies and a Description of the Hematopoietic Blast Colony Forming Assay

UNIT 1D.3

Elizabeth S. Ng,1 Richard P. Davis,1 Tanya Hatzistavrou,1 Edouard G. Stanley,1 and Andrew G. Elefanty1 1 Monash Immunology and Stem Cell Laboratories, Monash University, Clayton, Victoria, Australia

ABSTRACT This unit describes a protocol for the differentiation of human embryonic stem cells (hESCs). To generate spin embryoid bodies (EBs), known numbers of hESCs are deposited into low-attachment, round-bottomed 96-well plates in a serum-free medium supplemented with growth factors. The cells are then aggregated by centrifugation, initiating formation of EBs of uniform size. The spin EBs generated using this technique differentiate efficiently and synchronously along the lineages preferentially induced by the combinations of growth factors to which the cells are exposed. The 96-well format permits an assessment of the effects of different combinations of growth factors in the same experiment, facilitating the optimization of differentiation conditions for any given cell type. Up to 40 plates can be set up within a couple of hours by one experimenter, and aliquots of the differentiating EBs can be harvested at intervals and subjected to analyses C 2008 using a variety of techniques. Curr. Protoc. Stem Cell Biol. 4:1D.3.1-1D.3.23.
by John Wiley & Sons, Inc. Keywords: human embryonic stem cells r hESCs r differentiation r serum-free differentiation r spin EBs

INTRODUCTION This unit describes the differentiation of human embryonic stem cells (hESCs) as spin embryoid bodies (EBs) in a serum-free medium starting with enzymatically expanded stocks of hESCs. The first step is the propagation and expansion of human embryonic stem cells in bulk culture prior to differentiation (see Support Protocol 1 and Fig. 1D.3.1). Next, in Basic Protocol 1, spin EBs are generated by depositing known numbers of hESCs in round-bottomed 96-well plates in a serum-free medium supplemented with growth factors. The cells are then aggregated by centrifugation, initiating the formation of EBs of uniform size. Further differentiation of EBs is described in Basic Protocol 2, and harvesting EBs for further analysis is described in Support Protocol 2. The methylcellulose assay (Support Protocol 3) is used to detect clonogenic hematopoietic progenitors. The spin EBs generated using this technique differentiate efficiently and synchronously along the lineages preferentially induced by the combinations of growth factors to which the cells are exposed (Ng et al., 2005b; Pick et al., 2007; Costa et al., 2007; Davis et al., 2007). The 96-well format permits an assessment of the effects of different combinations of growth factors in the same experiment, facilitating the optimization of differentiation conditions for any given cell type.

Current Protocols in Stem Cell Biology 1D.3.1-1D.3.23 Published online January 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01d03s4 C 2008 John Wiley & Sons, Inc. Copyright


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The method is flexible and versatile, since up to 40 plates can be set up in a short time by one experimenter, and aliquots of the differentiating EBs can be harvested at intervals and subjected to analyses using a variety of techniques such as flow cytometry, further in vitro cultures, hematopoietic colony forming cell assays, transplantation, and interrogation of extracted RNA for gene expression using PCR or microarray platforms. Reproducible, scaleable, synchronous differentiation of hESC as spin EBs has many applications. Spin EB differentiations allow the researcher to access previously inaccessible populations of cells that emerge during early human development and provide insights into early human developmental biology. Spin EBs may be used in assay systems such as drug discovery platforms and directed differentiation to specific cell types for cell replacement therapies. NOTE: The following protocols are performed in either a Class I (laminar-flow) biosafety cabinet or a Class II biohazard hood. NOTE: All materials and reagents that come into contact with live cells must be sterile and proper aseptic technique must be used when handling the cells or setting up experiments. NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator unless otherwise specified. NOTE: It is important to stress that the successful formation of spin EBs requires the use of a serum-free differentiation medium and low-attachment round-bottomed 96-well plates. NOTE: The successful formation of spin EBs (described in Basic Protocol 1) is critically dependent on the prior enzymatic expansion of hESCs (described in Support Protocol 1). The transition to these larger-scale enzymatically passaged cultures from hESC stock cultures that are mechanically passaged to maintain genetic integrity is described in detail in UNIT 1C.1. BASIC PROTOCOL 1

GENERATION OF SPIN EMBRYOID BODIES (STAGE 1) Differentiation of the hESC may be considered as a two-stage procedure. In the first stage, spin EBs are formed in low attachment wells and the EBs grow and differentiate in response to exogenously added cytokines. During this phase of differentiation, hESC recapitulate early stages of post-implantation embryonic development and, in response to mesoderm- and endoderm-inducing stimuli, transiently express genes associated with the embryonic primitive streak that forms during gastrulation (Ng et al., 2005b; Costa et al., 2007; Davis et al., 2007). Following this stage, the differentiating spin EBs express genes marking mesoderm or endoderm germ layers. The induction of ectodermal lineages, typically neurons and glia, does not involve passage through this primitive streak intermediate stage. Spin EBs may be harvested at any time during this phase of the differentiation process and analyzed, or may be left to differentiate further (typically for a total of ∼10 days) prior to being transferred to flat-bottomed 96-well tissue culture plates for a second stage of differentiation (Basic Protocol 2). This second stage of adherent differentiation may last up to 2 to 3 weeks. Therefore, the total differentiation period (stages 1 and 2) may last for 25 to 30 days.

Materials

Directed Differentiation of hESCs as Spin Embryoid Bodies

hESCs in 75-cm2 or 150-cm2 tissue culture flasks at enzymatic passages between 5 and 20 (see Support Protocol 1) in hESC medium (see recipe for medium) 75- or 150-cm2 gelatinized (see recipe for gelatization of flasks) tissue culture flasks seeded with mitotically inactivated MEF at 2.0 × 104 /cm2 for maintenance of hESC or 0.8–1.2 × 104 /cm2 for passaging hESC prior to differentiation

1D.3.2 Supplement 4

Current Protocols in Stem Cell Biology

Sterile water Serum-free differentiation medium (see recipe) Cytokines required for differentiation (see recipes) CMF-PBS (Invitrogen) TrypLE Select cell dissociation enzyme (Invitrogen; see recipe) 0.4% Trypan blue (Fluka) 96-well round-bottomed low-attachment plates and lids (Nunc cat. no. 262162 and lids cat. no. 264122; or Costar cat. no. 3788) 14-ml sterile centrifuge tubes Tissue culture microscope with phase contrast objectives and phase rings Hemacytometer (Neubauer) Reagent reservoir, optional Multichannel pipets (Finnpipettes, Thermo Electron), optional 96-well plate spinner attachment for tissue culture centrifuge Set up spin EBs (day 0 minus 1) 1. On the day before differentiation (d0 minus 1), enzymatically passage hESCs (as described in Support Protocol 1) for at least two passages into two 150-cm2 flasks such that they will be 70% to 90% confluent on the day of set up (d0). This step is critical to the success of the differentiation. It ensures that the cells are actively dividing and removes dead and dying cells. The passaging ratio used to achieve this outcome depends on the bulk passage number of the cells and when they were last passaged. For example, hESCs that are at least passage 6 to 8 in bulk culture may require splitting 1:2 or 1:3 on the day prior to differentiation. Conversely, hESCs at passage 3 to 5 in bulk culture may simply require passaging to fresh feeders (denoted as a split of 1:1) and, if the cells are growing slowly or are very early in bulk culture, it may be necessary to pool the cells from two flasks into one fresh flask (denoted as a split of 2:1). The cells are passaged onto MEFs seeded at low density (0.8–1.2 × 104 /cm2 ) to effect a partial depletion of the feeder cells. Later-passage bulk cultures of hESCs may also be effectively passaged at 2:1 onto gelatin-coated flasks overnight as a feeder depletion step, but the recovery of cells on the day of differentiation has been found to be somewhat variable, therefore, this approach is not recommended. Similarly, cells may be passaged onto Matrigel as a substrate on the day prior to differentiation but this is expensive and labor intensive (in preparing the Matrigel). To calculate the number of hESC flasks to be passaged to provide cells to set up the differentiation on the following day, assume that sufficient undifferentiated hESCs will be obtained from one 150-cm2 or two 75-cm2 flasks to set up ∼20 96-well plates of spin EBs seeded at 3000 cells per well. Routinely, 5–6 × 106 cells are obtained from one 150-cm2 flask. Therefore, the yield from two such flasks (∼10–12 × 106 cells) will seed ∼40 plates (40 × 60 = 1800 wells at 3000 cells per well; requiring a total of ∼7.2 × 106 cells) leaving 3–5 × 106 cells available for replating onto a fresh 75-cm2 flask seeded with MEFs to maintain the undifferentiated culture. At the time of passaging onto low-density MEFs, an aliquot of cells may be taken for analysis of stem cell marker expression by flow cytometry to ensure that high-quality cells will be available for differentiation. Alternatively, this assessment can be done on cells left over when the differentiation experiment has been set up the following day (i.e., at day 0).

2. Prepare the required number of 96-well round-bottomed low-attachment plates. Round-bottomed low-attachment plates must be used or the spin EBs will not form. If nonsterile, molecular biology–grade plates are used, they require exposure to UV for at least 30 min. Plates and lids are placed faced up in a closed tissue culture hood under the UV light to sterilize. For convenience, the plates and lids may be left to sterilize overnight

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(but not longer, since the plastic is adversely affected with prolonged exposure to UV light). Once sterilized, replace lids on the plates and store in a sealed bag or container until use.

3. Add 80 µl sterile water to the 36 outer wells of each sterilized 96-well plate to prevent desiccation of the EBs during the differentiation time. The prepared plates may be stored in a sealed bag at room temperature or in the incubator prior to use the following day.

Form spin EBs: d0 of differentiation (stage 1) 4. Prepare differentiation medium and thaw aliquots of the cytokines required for the desired differentiation induction. Always check to see that there are enough cells and that they are in good condition for differentiation before making up differentiation medium. Do not add the cytokines to the medium (keep at 4◦ C) until the cells are harvested and counted and the exact volume of differentiation medium required can be calculated. While the growth factors chosen to initiate differentiation depend on the outcome desired by the researcher, they will commonly include those known to induce mesoderm, endoderm, and ectoderm in the developing embryo. For example, in the authors’ laboratory, BMP4 (20 to 40 ng/ml), VEGF (20 to 50 ng/ml), and SCF (40 ng/ml) are used to induce hematopoietic mesoderm differentiation and FGF2 (100 ng/ml) to induce ectoderm.

5. Harvest the hESC, which were passaged the day before onto low-density MEFs. Aspirate hESC medium and rinse once with 3 ml CMF-PBS to remove traces of medium. 6. Add 3 ml TrypLE Select per 150-cm2 flask and gently tilt the flask to coat the surface of the cells. Incubate 4 min at 37◦ C 7. Strike the base of the flask sharply one or two times on the bench surface of the culture hood to lift the cells. If the cells have not started to dislodge after 4 min, the flask may be returned to the incubator for an additional 1 min. Strike the flask again to dislodge the cells. 8. Add 5 ml CMF-PBS to the flask and collect the cells into a 14-ml sterile centrifuge tube. 9. Centrifuge cells 3 min at 480 × g, 4◦ C, and remove supernatant. Repeat the rinse step to ensure that all traces of dissociating enzyme are removed. 10. Resuspend the cell pellet in 5 ml of differentiation medium without growth factors or CMF-PBS per 150-cm2 flask of harvested hESCs. 11. Determine the cell concentration using a 10-µl aliquot of cells and hemacytometer to count a 1:1 dilution of cells with 0.4% Trypan blue. Remember there will be a residual amount of MEF contamination. As these cells are mitotically inactivated and adherence dependent, they will disappear during EB formation, but their initial presence may introduce a slight overestimation of the hESC count (<10%). Because the MEFs differ slightly in appearance under the hemacytometer from the hESCs, a skilled observer can actually distinguish between cell types.

12. Determine the total number of cells needed and the total volume of differentiation medium required for the experiment.

Directed Differentiation of hESCs as Spin Embryoid Bodies

Generally, 3000 hESCs in 100 µl of differentiation medium per well ensures reproducible EB formation for all the hESC lines that the authors’ laboratory has worked with.

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Figure 1D.3.1 hESC expanded in bulk culture prior to differentiation form colonies of uniform morphology and retain stem cell marker expression when analyzed by flow cytometry. (A to F) Bright field (A, D; BF) and fluorescence images (B, E; GFP) of the HES3 subclone, Envy, in bulk culture at 50× (A to C) and 100× (D to F) magnification 2 days after passaging using TrypLE Select. The merged bright field and fluorescence images (C, F; overlay) enable an unambiguous distinction between Envy colonies and feeder cells. (G and H) Flow cytometry profiles of Envy and MEL2 hESC lines stained with antibodies to the indicated stem cell markers (E-cadherin, KDR, GCTM2, CD9, CD117, and OCT4) demonstrating that they retain an undifferentiated phenotype when expanded in bulk culture. In all cases, 90% to 100% of the hESC express the indicated stem cell marker protein. Embryonic and Extraembryonic Stem Cells

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13. Add the cells to the medium. Calculate the volumes of cytokine required and add this to the medium containing the cells. Swirl gently to mix. When forming spin EBs for the first time with a new cell line, it is useful to perform an input cell titration experiment in which the number of cells used to generate the spin EBs varies across a range from 500 cells/well to 10,000 cells/well. The authors have observed that there is an optimum input number, often in the vicinity of 3000 cells/well.

14. Dispense 100-µl aliquots of cells in differentiation medium to each of the inner wells of the pre-prepared 96-well plates. To facilitate this process, the cells in differentiation medium can be added to a sterile reagent reservoir (Fig. 1D.3.2A) and 6 or 10 wells can be dispensed simultaneously using a multichannel pipet (Fig. 1D.3.2B). Always prepare ∼10% extra of cell stock in differentiation medium to ensure there are enough cells in differentiation medium for the requisite number of 96-well plates. Any leftover cells may also be dispensed into aliquots to form extra spin EBs.

Directed Differentiation of hESCs as Spin Embryoid Bodies

Figure 1D.3.2 Photographs of items of tissue culture equipment utilized during formation of spin EBs from hESC and their subsequent transfer to adherent culture. See text for more details. (A) A sterile, reusable reagent reservoir (with lid) used for dispensing hESC into low-attachment 96well plates for spin EB formation. (B) Multichannel pipettor with sterile tips attached for transferring cells from reagent reservoir into 96-well plate for spin EB formation. Also used for adding fresh medium to EBs prior to transfer to adherent wells and for affecting the actual transfer of EBs. (C) Plate spinner centrifuge attachment containing two 96-well plates used to aggregate hESC. (D) A sterile tip attached to vacuum tubing to allow aspiration of medium from the spin EBs. (E) A sterile Pasteur pipet placed in a pipet-aid used for collection of later stage EBs. Approximately 6 to 10 EBs may be collected at a time.

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Current Protocols in Stem Cell Biology

Figure 1D.3.3 Spin EBs formed and grown in the presence of BMP4, VEGF, and SCF generate hematopoietic mesoderm. (A) Bright field images of MIXL1GFP/w hESC aggregated at d0 (50×), which form discrete EBs by day 1 (100×). The EBs grow in size over the ensuing days of differentiation (day 2 to day 4, magnification 100×). (B) Bright field images of blast colonies derived from cells dissociated from day 4 spin EBs (d4EB) generated in the presence of BMP4, VEGF, and SCF and plated into methylcellulose supplemented with hematopoietic cytokines (SCF, VEGF, IL-6, IL-3, Tpo, and Epo). Images of colonies were taken after 3 to 16 days culture in methylcellulose (d3MC to d16MC) as indicated. In the final panel a May Grunwald Giemsa stained cytocentrifuge preparation from a colony after 25 days in MC shows the predominance of nucleated yolk sac-type erythroid cells. (C) Panels demonstrate erythroid and myeloid colonies derived from day 7 EBs disaggregated and plated in MC for 11 days (d11MC) and erythroid and myeloid cells growing in a stage 2 culture 11 days after replating onto a flat-bottomed tissue culture–treated well in the presence of hematopoietic growth factors (d10 EB+11d).

Embryonic and Extraembryonic Stem Cells

1D.3.7 Current Protocols in Stem Cell Biology

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15. Aggregate the cells in the 96-well plates by centrifugation to facilitate the formation of the EBs. Carefully stack the plates two- or three-high in a plate spinner (Fig. 1D.3.2C) and centrifuge the plates 3 min at 480 × g, 4◦ C, to aggregate the cells at the bottom of the wells so they will form spin EBs. 16. Carefully remove the plates from the spinner attachments so as not to dislodge cells and place in the incubator until required. Check at least one plate from each spin batch under the microscope to ensure cells have formed a pellet at the bottom of each round-bottomed well. Occasionally, it is necessary to repeat the spin step. Spin EBs will form by day 1 of differentiation usually surrounded by varying amounts of cell debris or dead cells (Fig. 1D.3.3A).

17. Incubate cells at 37◦ C. Harvest wells (see Support Protocol 2) at the times designated by the experimental design, up to 10 days. BASIC PROTOCOL 2

EXTENDED DIFFERENTIATION OF SPIN EBs (STAGE 2) After day 10 to 12 of differentiation, the spin EBs may be transferred to gelatinized wells of a 96-well flat-bottomed, tissue culture–treated plate in fresh medium to continue to differentiate. The differentiation medium at the platedown stage usually contains a different combination of growth factors from those that induced the first stage of EB differentiation in the low-attachment plates, depending on the desired outcome. An example of a day 10 EB plated down for a further 11 days in medium containing hematopoietic growth factors (SCF, VEGF, IL-3, IL-6, Tpo, and Epo) with the resultant emergence of erythroid and myeloid cells is shown in the last panel of Figure 1D.3.3C.

Materials Sterile water 0.1% (w/v) gelatin (see recipe) Spin EBs plated in round-bottomed, low-attachment plates (see Basic Protocol 1) Differentiation medium (see recipe) with required stage 2 growth factors (see recipes) 96-well flat-bottomed, tissue culture–treated plates Flexible tubing attached to a vacuum source (e.g., VacSax, VacSax Ltd.) Stereomicroscope (optional) Sterile reagent reservoir Multichannel pipettors (Finnpipettes, Thermo Electron) Transfer spin EBs to adherent culture: d10 platedown 1. Add sterile water to the outer wells and gelatinize the 60 inner wells of the required number of 96-well flat-bottomed, tissue culture–treated plates by adding ∼80 µl of 0.1% gelatin solution per well for a minimum of 20 min. 2. Using a sterile 20-µl tip attached to flexible tubing and a vacuum source (e.g., attached to a VacSax), aspirate most of the medium (leave ∼20 µl) from the wells containing the spin EBs to be transferred (Fig. 1D.3.2D).

Directed Differentiation of hESCs as Spin Embryoid Bodies

Performing this manipulation in a well-lit tissue culture hood makes it possible to identify each EB as a small white mass in the base of each well. If the EBs are difficult to see with the naked eye, visualize the whole plate at low power under a stereomicroscope in the laminar flow hood while aspirating the excess medium, to ensure that the EBs are not inadvertently sucked up in the process.

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Current Protocols in Stem Cell Biology

3. Add a volume of fresh differentiation medium with the required stage 2 growth factors to a sterile reagent reservoir. Transfer 100 µl to each EB-containing well using a multichannel pipettor. 4. Aspirate the gelatin from the gelatinized wells. 5. Set the volume of the multichannel pipettor to ∼120 µl (to take up all of the medium) and transfer the EBs from the round-bottomed low-attachment plates to the gelatinized tissue culture plates. Because the footprint of both types of plates is identical, the EBs can be transferred in the same pattern as they were set up, which reduces the chance of error if different wells contain spin EBs generated under different growth factor conditions.

6. Place plates in the incubator and leave EBs for 24 to 48 hr to attach and begin spreading. An alternative method of plating down EBs involves the use of 48-, 24-, or 6-well plates and transferring 2 or 8 or 30 EBs into the correspondingly larger well. Generally, use ∼100 µl medium volume per EB.

PROPAGATION AND EXPANSION OF HUMAN EMBRYONIC STEM CELLS IN BULK CULTURE PRIOR TO DIFFERENTIATION

SUPPORT PROTOCOL 1

The protocol for expansion of hESCs from stock cultures maintained by mechanical passaging is described in detail in UNIT 1C.1. The current protocol commences with hESC cultures that have been enzymatically passaged at least two times and have therefore been scaled up to a 75-cm2 tissue culture flask (UNIY 1C.1). hESCs are expanded and maintained in bulk culture for several passages prior to differentiation to allow them to adapt to enzymatic passaging. The hESCs in flasks grow closely together in monolayer colonies (Fig. 1D.3.1A) that can be readily passaged with enzymes every 3 to 4 days and maintain good viability, enabling the cultures to be gradually expanded. hESCs in early enzymatic passages (less than about passage 5) can only be maintained or minimally expanded (1:1 or 1:2) with each passage. However, by about passage 8, they can usually be expanded 1:3 or 1:4 and, by about passage 10, the cells not only dissociate more easily with enzyme, but also may require regular passaging at a ratio of 1:5. The quality of the enzymatically passaged hESC cultures can be objectively assessed by flow cytometry for a range of cell surface stem cell markers including E-cadherin, GCTM2, Tra-1-60, Tra-1-81, SSEA-3, SSEA-4, CD9, KIT (CD117), KDR, and for intracellular expression of OCT-4. In general, >90% of the hESCs should express these markers in a homogeneous fashion (see Fig. 1D.3.1B for examples). hESCs that have been enzymatically passaged 5 to 20 times are suitable for initiating spin EB differentiation cultures. Once cultures have been enzymatically passaged >20 to 25 times, they are discarded and replaced by freshly expanded stocks to minimize the emergence of aneuploid clones. In the authors’ laboratory, TrypLE Select or recombinant trypsin (TrypZean) are the dissociating agents of choice.

Materials hESCs in 75-cm2 tissue culture flasks at enzymatic passage 2 or greater Phosphate-buffered saline without Ca2+ and Mg2+ (CMF-PBS), sterile 1× TrypZean (TZ solution; Sigma; see recipe) or 1× TrypLE Select (TS solution; Invitrogen; see recipe), sterile Soybean trypsin inhibitor (required if TrypZean is used; Sigma; see recipe), sterile hESC medium (see recipe), sterile

Embryonic and Extraembryonic Stem Cells

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75-cm2 or 150-cm2 gelatinized flasks (see recipe) seeded with mitotically inactivated MEFs at 2 × 104 MEFs/cm2 (mitotically inactivated primary mouse embryonic fibroblasts, MEFs; Conner, 2000; UNIT 1C.3) Tissue culture microscope with phase contrast objectives and phase rings 14-ml centrifuge tubes Cell scraper (optional), sterile Refrigerated centrifuge Passage and expand hESC enzymatically 1. Begin with a confluent 75-cm2 flask of hESCs that are at least at passage 2 in bulk culture. 2. Aspirate the hESC medium and add 3 ml CMF-PBS to the flask, rinsing the surface of the cells to remove residual medium; aspirate CMF-PBS. 3. Add 2 ml of 1× TS solution, and incubate 4 to 5 min at 37◦ C or add 2 ml of 1× TZ solution, and incubate 3 min at 37◦ C. 4. Strike the flask firmly one or two times on the base of the tissue culture hood to dislodge the hESCs. If the cells were dissociated using TZ, neutralize with 1 ml soybean trypsin inhibitor.

5. Add 5 ml hESC medium, mix, and transfer the cells to a sterile 14-ml centrifuge tube. If the majority of the cells do not detach when the flask is struck against the base of the tissue culture hood, a cell scraper may be required to dislodge them. This is preferable to leaving the cells in enzyme solution for a prolonged period and will not adversely affect their viability. This problem often arises if the cells are not passaged frequently enough.

6. Centrifuge cells 3 min at 480 × g, 4◦ C in a refrigerated centrifuge. While the cells are spinning, remove the medium from a 150-cm2 flask previously seeded with 2 × 104 MEFs/cm2 and add 20 ml fresh hESC medium. 7. Decant the supernatant from the centrifuged cells. Gently resuspend cell pellet in 5 ml of fresh hESC medium. Add cell suspension to the 150-cm2 flask, swirl to mix the cells, and place the flask in the incubator. The hESC reattach within a few hours and form many small, evenly sized colonies that will be ready to passage again in ∼3 to 4 days. SUPPORT PROTOCOL 2

HARVESTING EBs FOR FURTHER ANALYSIS Spin EBs may be harvested at any stage during differentiation, the number of EBs or plates of EBs harvested depends on the subsequent analysis to be performed. The method of harvesting varies with the developmental stage of the EBs and whether the harvested tissue must or must not be sterile.

Materials Stage 1 cultures (see Basic Protocol 1) or stage 2 cultures (see Basic Protocol 2) Appropriate lysis buffer for nucleic acid extraction, flow cytometry buffer, or differentiation medium Phosphate buffered saline without Ca2+ and Mg2+ (CMF-PBS) TrypLE Select (see recipe) Directed Differentiation of hESCs as Spin Embryoid Bodies

10-, 14-, or 50-ml centrifuge tubes, sterile P200 Gilson pipetman with sterile (plugged) 200-µl tips Dissecting microscope, optional

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Current Protocols in Stem Cell Biology

Multichannel pipettor Pasteur pipets and pipet-aid 23-G 1-in. needle 3-ml syringe For spin EBs from stage 1 cultures (d1 to d10) 1a. If sterile cells are required for further analyses, harvest the plates in the tissue culture hood. Conversely, if sterility is not required, EBs may be harvested from plates on the laboratory bench.

2a. Prepare labeled collection tubes (10-, 14-, or 50-ml sterile tubes) depending on the number of plates to be harvested. Calculate the approximate total volume of medium being harvested, based on an estimate of 6 ml per 96-well low-attachment plate. 3a. Harvest individual spin EBs differentiated for 1 to 5 days using a P200 Gilson pipetman with a sterile (plugged) 200-µl tip set at 120 µl. This can be done under direct vision and the wells checked afterwards under the dissecting microscope to ensure removal of the EBs. When collecting into a 10- or 14-ml tube, a multichannel pipettor may be used with two sterile tips to simultaneously harvest two spin EBs. If convenient, collect into a 50-ml tube because the tube diameter is large enough to accommodate three tips attached to the multichannel pipettor, thus allowing three EBs to be simultaneously harvested. If multiple plates are to be harvested, the collection tubes may be placed on ice during the procedure to maximize cell viability. This method of collection is desirable for early stage spin EBs, which are easily damaged by rough handling. Use a more rapid method to harvest EBs at d6 to d10, which are larger and more robust. Place a sterile unplugged Pasteur pipet into a pipet-aid (Fig. 1D.3.2E) and by gentle control of the suction, collect spin EBs sequentially under direct vision, from ∼6 wells and then dispense them gently into an appropriately labeled tube.

4a. Centrifuge EBs 3 min at 480 × g, 4◦ C and remove excess supernatant. If samples from several tubes are to be pooled, leave some residual medium in the tubes, dislodge the EBs, and transfer to a single tube using a 1-ml Gilson pipettor. Always transfer the EBs in residual medium rather than CMF-PBS, to prevent the EBs from adhering to the internal surface of the plastic tip.

5a. Place the harvested EBs directly into an appropriate lysis buffer for nucleic acid extraction or dissociate into single cells for flow cytometric evaluation or colony forming cell assays (see Support Protocol 3).

For spin EBs from stage 2 cultures (>d10) Harvesting EBs from stage 2 cultures that have been plated down (i.e., after day 10) requires a slightly modified collection procedure. 1b. If some of the differentiated cells are in suspension (as observed, for example, with day 15 to 20 EBs generated in the presence of hematopoietic growth factors), first collect the medium from each well using a multichannel pipettor into a 50-ml tube. 2b. Rinse each well of the plate with 80 µl of CMF-PBS. Add the CMF-PBS wash to the 50-ml tube to ensure that cells are not lost. Keep the collection tube on ice while the EBs are disaggregated as described below.

Embryonic and Extraembryonic Stem Cells

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3b. Add 80 µl of TrypLE Select to each well and place 10 to 30 min in a 37◦ C incubator, depending on the size of the EBs. For example, day 12 EBs usually require only a 10- to 15-min treatment while EBs older than day 15 often require 20 to 30 min in enzyme. This enzymatic treatment will lift and dissociate the adherent cells that have grown out from the EB mass following platedown. The main mass of the EB will be partially dissociated and the EB will lift easily from the well.

4b. Dislodge each EB with a pipet tip and collect the partly disaggregated EBs into a separate tube. 5b. Complete the disaggregation of the EBs by passing them one to two times through a 23-G needle attached to a 3-ml syringe. Older, larger EBs may need to be passed first through a larger-gauge needle (e.g., 21-G) before completing the disaggregation with a 23-G needle.

6b. Pool the harvested suspension and adherent fractions into a single tube and centrifuge 3 min at 480 × g, 4◦ C. 7b. Transfer cell pellet to a 10-ml tube and repeat wash step (6b) with an additional 5 ml CMF-PBS to ensure complete removal of the TrypLE Select. 8b. Place cell pellet in an appropriate lysis buffer for nucleic acid extraction, resuspend in an appropriate buffer for flow cytometry (remember to filter the cell suspension first), or resuspend in a small volume of differentiation medium for colony forming cell analysis (see Support Protocol 3). SUPPORT PROTOCOL 3

IDENTIFICATION OF HEMATOPOIETIC PROGENITOR CELLS IN SPIN EBs USING A METHYLCELLULOSE COLONY FORMING ASSAY Spin EBs may be harvested during differentiation and hematopoietic progenitors identified by their ability to form colonies in methylcellulose. Hematopoietic colony forming cells can be readily identified in spin EBs generated in the presence of BMP4, VEGF, and SCF from day 3 to 4 of differentiation onwards. The spin EBs are harvested, dissociated into single cells, and plated into low-attachment wells in methylcellulose, a semi-solid medium, supplemented with hematopoietic cytokines. The number and type of hematopoietic colonies observed reflect the frequency and spectrum of progenitor cells present in the differentiated spin EB cell population. The earliest progenitors, observed at day 3 to 4, are called blast colonies, and arise from a multi-potential precursor (the blast colony forming cell, Bl-CFC) that typically generates large numbers of primitive, yolk-sac-type nucleated erythroid cells, myeloid cells, and, in many cases, endothelial cells (Costa et al., 2007; Kennedy et al., 2007; Davis et al., 2007). Spin EBs differentiated in a combination of BMP4, VEGF, and SCF upregulate the primitive streak markers MIXL1 and BRACHYURY by day 3, indicative of the induction and patterning of mesodermal cells within the EBs. One of the earliest types of mesodermal cells specified is the yolk-sac-type blast colony forming cell (Bl-CFC), loosely termed the hemangioblast, which is capable of clonally differentiating into primitive erythroid, myeloid, and adherent (endothelial) cells when placed in a semi-solid colony forming assay. Bl-CFCs are generally found in day 3 to 5 spin EBs, although, in the authors’ laboratory, the peak number of Bl-CFCs has been observed in day 4 spin EBs for a number of independent hESC lines.

Directed Differentiation of hESCs as Spin Embryoid Bodies

1D.3.12 Supplement 4

Current Protocols in Stem Cell Biology

Materials Spin EBs at day 4 of differentiation (at least 30 to 60 EBs required) CMF-PBS (Invitrogen) TrypLE Select (Invitrogen; see recipe) Differentiation medium (see recipe) 0.4% Trypan blue (Fluka) MethoCult, serum-free (MC, Stem cell Technologies; see recipe) Hematopoietic cytokines (PeproTech, see recipes) Sterile water Sterile 10-ml polypropylene yellow-cap tubes 37◦ C water bath 23-G, 1-in. and 18-G, 1 1/2-in. needles (Terumo) 3-ml syringe (Terumo) Sterile FACS tubes (5-ml polystyrene, 12 × 75–mm) with lids and with cell-strainer caps (BD Falcon) Nescofilm (Bando Chemical Ind. Ltd) or equivalent Hemacytometer 24-well low-attachment plates (Nunc cat. no. 144530) Gilson pipettors (John Morris Scientific) Harvest and dissociate spin EBs at day 4 1. Collect at least 30 day 4 spin EBs into a 10-ml yellow-cap polypropylene tube. Change the tips between collections if collecting spin EBs from more than one set of differentiation conditions. If collecting more than 60 EBs (more than 6 ml), collect into a 14- or 50-ml tube. Collection is more rapid using a multichannel pipettor with two 200-µl sterile plugged tips attached (volume set at 120 µl) to collect the spin EBs two at a time (see above).

2. To pellet the EBs, briefly centrifuge 1 min at 480 × g, 4◦ C. Aspirate most of the medium, leaving ∼500 µl. Transfer the EBs in this residual medium to a 10-ml tube. Transfer the EBs in residual culture medium rather than in CMF-PBS to prevent them from sticking to the plastic pipet tip. Transferring to a 10-ml tube facilitates the subsequent dissociation step.

3. Wash the EBs with 5 ml CMF-PBS to remove residual differentiation medium and centrifuge EBs 3 min at 480 × g, 4◦ C. 4. Aspirate CMF-PBS, tap the tube to dislodge the EBs in the residual ∼50 µl CMFPBS remaining in the tube, and add 1 ml of TrypLE Select. Place tubes 8 to 12 min in a 37◦ C water bath. The dislodged EBs should float briefly in the TrypLE solution. The duration of incubation in enzyme depends on the size of the EBs. At day 4 of differentiation, even large EBs will dissociate easily in TrypLE Select in 8 to 12 min.

5. After incubation in TrypLE Select, remove the tubes from the water bath and place them in a rack in the tissue culture hood. Remove all the caps and, one sample at a time, dissociate the cells by passing the EBs one to three times through a 23-G needle attached to a 3-ml syringe. Perform this step gently and without too much force, to prevent lysing the cells. As the EBs break apart and dissociate into single cells, the solution becomes cloudy. If dissociating more than one sample of day 4 EBs, prepare the needles and syringes in advance. Embryonic and Extraembryonic Stem Cells

1D.3.13 Current Protocols in Stem Cell Biology

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Wash dissociated cells 6. Add 2 ml of differentiation medium without cytokines to each tube and top up with 5 ml CMF-PBS to dilute the TrypLE Select. Centrifuge cells 3 min at 480 × g, 4◦ C and aspirate the supernatant. The cells initially pellet more easily when resuspended in a mixture of medium and CMF-PBS.

7. Repeat wash step (step 6) with an additional 5 ml CMF-PBS to ensure complete removal of the TrypLE Select. 8. Resuspend the dissociated cells from 30 EBs in 200 µl of differentiation medium without cytokines. Filter cells using a sterile FACS tube with a cell-strainer cap. To reduce the possibility of contamination, open the packet of cell-strainer cap tubes in the tissue culture hood. Place the required number of tubes in a rack. Gently add the dissociated cells to each cap and cover the caps with a small square of Nescofilm. If more than 30 EBs have been dissociated, the cell pellet will be larger. Resuspend the cells in a proportionately greater volume than 200 µl. The object of this is to enable an accurate cell count, i.e., not too many or too few cells in the 10-µl volume that will be taken to perform the count.

9. Briefly centrifuge cells 1 min at 480 × g, 4◦ C, to collect the cells into the sterile FACS tube. This step removes cell clumps and cellular debris and ensures that the cells are in a single-cell suspension.

Count cells in samples 10. Place tubes in a rack in the tissue culture hood. Remove Nescofilm and the cellstrainer caps. Gently resuspend the cells in the medium and remove 10 µl to perform a cell count using a hemacytometer. If there are multiple samples to count, place 10-µl volumes of 0.4% Trypan blue in wells of a Terasaki or 96-well plate, arranged in columns. In adjacent wells, place 10 µl of each sample to be counted.

11. Use a separate sterile tip for each sample. Recap the tubes with caps taken from sterile capped FACS tubes to prevent desiccation and to keep the samples sterile. Store the capped sample tubes on ice or at 4◦ C until required. The remaining uncapped tubes can be used for any future non-sterile FACS analyses.

12. Perform a cell count. In the case of multiple samples, take a 10-µl droplet of 0.4% Trypan blue and add it to the 10-µl cell droplet in the adjacent well and mix gently before loading the hemacytometer. Mixing the dye and cells immediately prior to counting reduces the detrimental effects on cell viability of prolonged exposure of the cells to Trypan blue.

13. Determine the total cell number for each sample and calculate the volume that will deliver a total of 30,000 to 45,000 cells to be spread across three replicate wells at 10,000 to 15,000 cells per well. These input cell numbers have been chosen as they usually contain progenitors for at least 100 blast colonies. If there are too many or too few Bl-CFCs, the input number can be adjusted appropriately.

Directed Differentiation of hESCs as Spin Embryoid Bodies

In general, 30 EBs at day 4 will yield 0.35–0.5 × 106 cells, which is in excess of the number required for the colony forming assay. The remaining cells may be recultured, used to make RNA, or analyzed by FACS for cell surface marker expression.

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Current Protocols in Stem Cell Biology

Prepare methylcellulose wells 14. Prepare the MethoCult (MC) stock. The total volume of stock required is determined by the total volume of the triplicate wells to be set up plus an additional ∼20% to take into consideration the difficulty in dispensing the viscous reagent. For example, for three wells of 400 µl methylcellulose, dispense 1.5 ml of MC into a 10-ml tube. Add cytokines to the tube of MC. Shake vigorously to mix and centrifuge the tube briefly to collect the MC and remove the air bubbles caused by the mixing. Cytokines are added to MC at a final concentration of: stem cell factor (SCF) 100 ng/ml, vascular endothelial growth factor (VEGF) 50 ng/ml, interleukin (IL)-3 30 ng/ml, interleukin (IL)-6 30 ng/ml, thrombopoietin (Tpo) 30 ng/ml, and erythropoietin (Epo) 3 U/ml.

15. Label one 10-ml tube for each triplicate MC sample. Add 30,000 to 45,000 cells in a volume of ∼100 to 150 µl to each tube. Use an 18-G needle attached to a 3-ml syringe to add 1.5 ml of MC to each tube. Use a separate needle and syringe for each sample. 16. Mix the cells into the MC by drawing the cells and MC gently into and out of the syringe three to five times (taking care not to generate too many air bubbles). 17. Finally, draw the cells and MC into the barrel of the syringe. Let the syringe sit for 1 min in the tube to allow the residual MC to collect in the base of the tube and draw this into the syringe as well. 18. Dispense the MC mix into three wells of a 24-well low-attachment plate. Use a 20-µl Gilson pipettor to remove any large air bubbles from the MC. 19. Add sterile water to the outer wells of the plate to reduce desiccation of the MC during the ensuing 10- to 15-day culture period. Place the plate in the incubator and leave undisturbed for 5 days. For improved hemoglobinization of the blast colonies, place the cultures in a low-oxygen environment (5%O2 /5%CO2 /90%N2 ) at 37◦ C in a humidified incubator.

20. Check for blast colonies formation during the first 5 to 7 days in MC. They are visibly hemoglobinized after ∼9 days and are optimally scored from day 10 to day 16 (Fig. 1D.3.3B).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Cytokines Recombinant human erythropoietin (PeproTech) Reconstitute lyophilized powder in distilled water to 0.1 to 1 mg/ml. Dilute to 1 U/µl in CMF-PBS and dispense into 50-µl aliquots and store at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human vascular endothelial growth factor (VEGF165 , PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in distilled water. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C.

continued

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Recombinant human stem cell factor (PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in 10 mM acetic acid. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human interleukin-3 (PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in distilled water. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human interleukin-6 (PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in distilled water. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human thrombopoietin (PeproTech) Reconstitute lyophilized powder at 0.5 to 1 mg/ml in 5 to 10 mM Tris·Cl, pH 7.6. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C.

Gelatin, 0.1% (w/v) Add 0.5g of gelatin powder (from porcine skin; Sigma cat. no. G1890) to 500 ml distilled water and autoclave to dissolve and sterilize. Store up to 6 months at room temperature.

Gelatinization of plates and flasks Prior to addition of MEFs, add enough 0.1% gelatin solution (see recipe) to cover the base of all plates/flasks. Let stand for 10 to 15 min to coat the surface and remove by aspiration immediately prior to addition of MEFs.

Growth factors for differentiation Recombinant human bone morphogenetic protein 4 (BMP4, R&D Systems) Resuspend lyophilized powder at 100 ng/µl in 4 mM HCl and store as 10- to 30-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human vascular endothelial growth factor (VEGF, PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in distilled water. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C. Recombinant human stem cell factor (SCF, PeproTech) Reconstitute lyophilized powder at 0.1 to 1 mg/ml in 10 mM acetic acid. Dilute to 50 ng/µl in CMF-PBS and store as 50-µl aliquots at −80◦ C. Store thawed aliquots for 1 to 2 weeks at 4◦ C.

hESC medium

Directed Differentiation of hESCs as Spin Embryoid Bodies

DMEM/F12 (Invitrogen) containing: 20% (v/v) knockout serum replacer (Invitrogen) 10 mM non-essential amino acids 2 mM L-glutamine (or GlutaMaxI) 50 mM 2-mercaptoethanol 1× penicillin/streptomycin (200× stock from Invitrogen) continued

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10 ng/ml FGF2 (see recipe) Filter sterilize using a 0.22-µm Stericup filtration unit (Millipore). Store up to 1 week at 4◦ C. hESC differentiation medium (BPEL) The base medium is Iscove’s modified Dulbecco’s medium (IMDM, phenol red–free) mixed with Ham’s F12 nutrient mixture in a 1:1 ratio. Additional components are: 0.25% (w/v) final deionized bovine serum albumin (BSA, Sigma) A 10% stock solution of BSA in water is deionized through three changes of resin beads (AG501-XB(D); 20 to 50 mesh; BioRad cat. no. 142-6425) prior to use.

0.25% (w/v) final polyvinyl alcohol (PVA, Sigma) The 5% or 10% PVA stock solution is usually prepared by dissolving the solid in distilled water for several days at 4◦ C. Alternatively, to facilitate initial solubilization, the PVA solution can be incubated overnight at 37◦ C or heated 1 to 2 min to 85◦ to 90◦ C. The PVA solutions may be pre-filtered through a 0.45-µm filter unit (INTERPATH PES filter, Nylon membrane) to reduce viscosity.

αMono-thioglycerol (350 to 450 µM; Sigma) 0.05 mg/ml final ascorbic acid 2-phosphate/ascorbic acid (Sigma) 2 mM GlutaMaxI (Invitrogen) 5% (v/v) final PFHMII (protein-free hybridoma medium, Invitrogen) Linoleic acid (100 ng/ml final; Sigma) Linolenic acid (100 ng/ml final; Sigma) SyntheChol (2.2 µg/ml final; Sigma) Insulin-transferrin-selenium (100× stock ITS-X, Invitrogen) 1× penicillin/streptomycin (200× stock, Invitrogen) MEF medium DMEM (4.5 g/liter glucose, without L-glutamine and sodium pyruvate, Invitrogen) containing: 10% (v/v) batch-tested, heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine (Invitrogen) 1× penicillin/streptomycin (200× stock, Invitrogen) Filter sterilize and store up to 2 weeks at 4◦ C MethoCult MethoCult, serum free, without cytokines (Stem Cell Technologies cat. no. 04236), stored at −20◦ C. Thaw the contents (80 ml) of the stock bottle at 37◦ C. Shake the contents vigorously to ensure the MethoCult is well mixed. Let the contents of the bottle settle (the air bubbles will dissipate) and label sterile 50-ml tubes with details of the lot number and other information to help keep track of batch-to-batch variation. Use a 20-ml syringe (without a needle attached) to dispense 10- or 20-ml aliquots into the labeled 50-ml tubes. Use a fresh syringe with an 18-G needle and then a 1-ml tip on a Gilson pipettor to collect the last 10 ml or so from the stock bottle. The solution is very viscous, therefore, <80 ml will be recovered from the stock bottle. An 80-ml bottle will usually give three 20-ml and one 15- to 17-ml aliquots. Store aliquots at −20◦ C. To use: Thaw MethoCult (overnight at 4◦ C or for a few minutes in a 37◦ C water bath), add the appropriate cytokines, and shake tube to mix. Briefly centrifuge (20 to 30 sec) to collect the viscous mixture and reduce the air bubbles. Current Protocols in Stem Cell Biology

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Recombinant basic human fibroblast growth factor (bFGF/FGF2) Resuspend lyophilized bFGF (PeproTech) to a final stock concentration of 10 ng/µl (10 µg/ml; 1000×) in CMF-PBS containing 0.1% (w/v) BSA or recombinant human albumin and 1 mM DTT. Dispense into 100- to 500-µl aliquots and store at −80◦ C.

Soybean trypsin inhibitor Soybean trypsin inhibitor (STI, Sigma) is used to neutralize either trypsin or TrypZean. Weigh the powdered stock (Sigma) and prepare a 1 mg/ml solution in CMF-PBS. Filter sterilize and dispense into 5- or 10-ml aliquots and store up to 6 to 12 months at −20◦ C. Thawed aliquots are stable stored for 1 to 2 weeks at 4◦ C. To use, add a volume equal to half that of the trypsin or TrypZean solution used to dissociate cells and swirl to mix. Proceed to collect cells by centrifuging 3 min at 480 × g, 4◦ C.

TrypLE Select Aliquot 1× TrypLE Select (Invitrogen cat. no. 12563-029) stock solution into smaller volumes (20- to 50-ml) and store up to 4 months at 4◦ C.

TrypZean solution Thaw 1× TrypZean solution (Sigma cat. no. T 3449) stock solution and aliquot into smaller working volumes. Store up to 12 months at −20◦ C.

COMMENTARY Background Information

Directed Differentiation of hESCs as Spin Embryoid Bodies

Embryonic stem cells are pluripotent, continuously growing cells derived from the inner cell mass of the pre-implantation, blastocyst stage mammalian embryo (Evans and Kaufman, 1981; Martin, 1981; Thomson et al., 1998; Reubinoff et al., 2000). Mouse ESCs are able to contribute to cells of all lineages in the developing embryo and both mouse and human ESC form teratomas containing cellular derivatives of all three germ layers (Evans and Kaufman, 1981; Martin, 1981; Thomson et al., 1998; Reubinoff et al., 2000). Differentiating mouse and human ESC recapitulate aspects of early mammalian development in response to exogenously supplied growth factors. They sequentially express cohorts of genes that mark specific stages of postimplantation embryonic development, starting from the inner cell mass stage and passing through a series of developmental fate restrictions before expressing genes that mark derivatives of mesoderm, endoderm, and ectoderm germ layers (Keller et al., 1993; Robertson et al., 2000; Lacaud et al., 2002; Ng et al., 2005a,b; Hirst et al., 2006). This developmental concordance between embryonic development and in vitro ESC differentiation is reflected by the fact that the same growth factors that induce and pattern the embryo also allow directed differentiation of ESC (Smith et al., 2007; UNIT 1D.1).

To capitalize on the potential research and regenerative medicine possibilities promised by embryonic stem cells, it will be necessary to reproducibly and robustly direct their differentiation along lineages of interest. Efficient differentiation of human ESCs is problematic, at least in part due to the fact that hESCs survive poorly as isolated single cells, the usual starting point for mouse ESC differentiation. Therefore, it was believed that a cell aggregate of some kind would be necessary to successfully initiate differentiation. When mechanical dissection or enzymatic generation was attempted on hESC colony pieces to initiate EB formation, an unacceptable variability and asynchrony in differentiation outcomes that made it very difficult to assess the effects of the growth factors used to direct differentiation were observed. The next approach that was tried was to create an EB using a known starting number of cells that would provide a critical mass of cells that would survive, proliferate, and differentiate in response to growth factors. The spin EB method that emerged has several advantages over prior hESC differentiation methods. Spin EBs are uniform in size and morphology and their differentiation is synchronous across the wells in a culture plate. As a consequence, it is possible to objectively assess the ability of exogenously added growth factors to direct differentiation when the spin EB method

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is combined with a serum-free differentiation medium. The method allows as many plates of EBs to be generated as necessary, so that the size of an experiment can be tailored to suit, for example, the numbers of growth factors to be assessed and type of analysis that will be performed. The availability of ESC lines in which the genetic loci of key developmentally regulated genes are molecularly tagged with reporter genes, such as green fluorescent protein (GFP), adds a further dimension to the utility of this method. For example, the human (and mouse) ESC lines carrying GFP inserted into the locus of the gastrulation marker MIXL1 facilitate the rapid objective assessment of growth factors that induce differentiation to the mesodermal and endodermal precursors of the primitive streak (Ng et al., 2005a; Costa et al., 2007).

Critical Parameters and Troubleshooting Spin EBs There are several critical points during the set up of spin EBs that influence the success or failure of the differentiation experiment. First, ensure that a homogenous population of undifferentiated stem cells is retained during the hESC expansion phase. For both the experienced scientist and the less-experienced researcher, regular analysis of stem cell markers by flow cytometry will provide an objective means to monitor the quality of the starting population. Care must also be taken to ensure maximum viability of the cells, especially at early passage in bulk culture as the cells are more fragile and susceptible to enzymatic damage during harvesting. It is important to be aware that the batch of mitotically inactivated MEFs significantly impacts on the general health and well-being of the hESC cultures. A bad batch of MEFs may cause the cells to grow more slowly so that the cells require a longer interval between successive expansion passages, or need to be expanded more cautiously. Bad MEFs may also adversely affect the percentage of cells that express stem cell markers. Changing to a fresh batch of MEFs usually solves this problem. If hESCs do not replate well during passaging/expansion steps, it may also be due to traces of TrypLE Select enzyme solution carried into the medium following dissociation of the cells. It is important to rinse the cell pellets well after dissociation.

Passaging of the expanded hESC cultures on the day prior to differentiation (day 0 minus 1) is also an important step in ensuring the optimum quality and yield of cells for differentiation. Cells in cultures that become over confluent may take on a “fried egg” appearance with more abundant cytoplasm than usual, probably indicating some early visceral endoderm differentiation. Conversely, cultures containing cells that are not confluent enough tend to differentiate poorly and make EBs that are smaller than expected for the number of cells seeded. When harvesting hESC for differentiation as spin EBs, ensure the cells are well rinsed to remove all traces of TrypLE Select as there is no specific inhibitor available to neutralize the solution. This is usually achieved by giving the cells an extra rinse in CMF-PBS before performing a cell count. Residual TrypLE Select will slowly kill the cells. After harvesting the cells, place them on ice to preserve their viability while a cell count is completed. At this stage the cells are resuspended in CMF-PBS, which is not a very supportive medium. Factors that may contribute to EB death or poor differentiation during the first 4 days include: (1) residual TrypLE Select in the differentiation medium; (2) an inaccurate cell count resulting in the formation of EBs from too few input cells and a subsequent failure to develop (EBs are small and often contain brown pigmented cells); (3) uneven mixing of the cells at the time of dispensing into the 96-well plates resulting in EBs that are of variable size, from too small to too large. Small EBs (e.g., <1000 cells) often fail to grow and tend to contain brown pigmented cells; very large EBs (>4000 cells) fail to differentiate well along mesodermal lineages (e.g., in response to BMP4) and tend to form a preponderance of neural cell types. (4) Inadequate concentration or weak activity of the batch of growth factors used to induce differentiation; factors such as BMP4 positively influence cell viability as well as induce differentiation during the early stages of spin EB differentiation. Individual batches of growth factors need to be titrated to determine their optimal concentration range for use. The authors have observed several-fold variation in activity between sequential, commercially obtained individual batches of growth factors. A balance between the cell number and concentration of inductive growth factor is important. For example, the combination of 30 ng/ml BMP4 in

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serum-free medium will induce mesoderm efficiently in spin EBs generated with 3000 cells per well. (5) Incorrect preparation of the differentiation medium: don’t leave anything out and specifically don’t add too much α-monothioglycerol (αMTG). Finally, never add serum to the spin EB differentiation medium for the first stage. It means instant failure because the cells will stick to the wells and spin EBs will not form. If it is absolutely required for a specific protocol, serum can be added during the second, adherent cell stage of the differentiation.

Directed Differentiation of hESCs as Spin Embryoid Bodies

Colony forming assay The number of hematopoietic blast colony forming cells at days 3 and 4 of differentiation is dependent on a number of factors including the cell line used, the differentiation medium and growth factor combination, and the number of cells added to each well of MC. The authors’ experiments suggest that too few input cells (1000 to 5000 cells/400 µl MC well, depending on the cell line) may result in a disproportionately low number of colony forming cells, suggesting that there may be an autocrine or paracrine component influencing colony frequency. Conversely, if plates are too crowded (over 400 colonies/well), related to a high clonogenic precursor frequency or to the use of too large an input cell number (>20,000 cells/well), the growth and maturation of erythroid colonies is inhibited by overcrowding. The authors find that 10,000 to 15,000 input cells per well give 100 to 300 Bl-CFCs per well of MC. A good number for the inexperienced user to start with is 15,000 cells per well. If there are >300 colonies per well, then the input number can be dropped back to 10,000 per well. Following scoring of the colonies after 10 to 16 days in MC, individual colonies may be plucked for morphological or PCR analysis or the contents of entire wells may be harvested, and analyzed by FACS or RNA extracted for gene expression studies. Colony forming assays may be performed using cells dissociated from EBs at other time points during differentiation after days 3 to 4 (Fig. 1D.3.3C). EBs at later stages of differentiation are much larger and the cell yield is potentially greater but they are more difficult to dissociate. To ensure that enough cells are acquired following dissociation, harvest 20 to 30 EBs. The initial stages of EB collection are the same as for day 4 EBs. The dissociation of later stage EBs requires longer incubation times in TrypLE Select. The outer cell layer

is particularly recalcitrant to dissociation and can be tough and stringy, possibly due to the extracellular matrix produced by the cells. For example, incubate day 6 to 7 EBs for 15 to 20 min and incubate day 10 EBs for 20 to 30 min. Large day 10 EBs may be removed from the 37◦ C water bath at the 20-min mark and passed once or twice through a 26-G × 1-in. needle attached to a 3-ml syringe to break open the outer cell layer before incubation for an additional 10 min at 37◦ C. With either variation, the outer cell layer may remain incompletely dissociated and may get very viscous, trapping other cells. Microscopic examination may reveal viable cells trapped onto strands of matrix-like material. Therefore, a filtration step using the FACS tubes with the cell-strainer caps is needed to produce a singlecell suspension. In cultures that are continued longer that 10 days, the spin EBs are usually transferred to flat-bottomed tissue culture–treated plates to encourage adherence of cells. The cultures flatten out somewhat under these conditions, and they will generally require a similar period of dissociation in TrypLE select as the day 10 spin EBs.

Anticipated Results Spin EBs begin to form within 1 day of differentiation (Fig. 1D.3.3A), usually surrounded by a variable amount of cellular debris. The EBs remain small during the first few days of differentiation. By day 2 to 3, an outer layer (perhaps representing visceral endoderm) can be observed around the periphery of the spin EB. There is a visible increase in size of the EBs from days 3 to 4 of differentiation. By day 4 of differentiation, the yield from 60 EBs (one 96well plate) after dissociation into single cells should be ∼0.7–1 × 106 cells. Gene expression analysis should confirm differentiation. For example, genes marking the advent of the primitive streak (gastrulation; e.g., MIXL1, brachyury, and goosecoid) should be transiently expressed for a few days from day 3 for cultures differentiated in the presence of BMP4 or activin (Fig. 1D.3.4). The authors have found that surface expression of the platelet-derived growth factor receptor α (PDGFRα), detected by flow cytometry of dissociated cells between day 3 and day 10, is a sensitive indicator for the emergence of mesoderm in response to BMP4 (Fig. 1D.3.5).

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Figure 1D.3.4 Gene expression profiles of HES3 spin EBs differentiated in BMP4, VEGF, and SCF analyzed at days 0, 2, 4, 6, 8, and 11 by real-time PCR. During differentiation, expression of the stem cell gene OCT4 is downregulated prior to upregulation of the primitive streak genes MIXL1 and brachyury. This is followed by expression of genes marking early hematopoietic mesoderm (GATA2, RUNX1, and CD34) and then genes marking cells committed to the erythroid lineage (GATA1 and γ-globulin). Expression of the target gene is shown normalized to GAPDH as a reference gene (relative gene expression) on a log scale on the y-axis.

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Figure 1D.3.5 Flow cytometric analysis of day 11 spin EBs generated from Envy cells differentiated in BMP4, VEGF, and SCF, demonstrating that the majority (∼67%) of cells express PDGFRα but very few cells (<2%) express CXCR4. This results indicates predominant differentiation to mesoderm in response to stimulation by these growth factors. Expression of PDGFRα is seen from day 3 onwards, peaks at approximately day 6 to 7 and is maintained at high levels until days 10 to 12.

Transfer of EBs to adherent culture allows growth and continued differentiation of adherent cell types and creates a slightly different interactive environment for cells differentiating in the EBs. For instance, under appropriate growth factor stimulation, foci of spontaneously beating cardiomyocytes will be evident ∼5 days after transfer.

Time Considerations It should take 1 to 2 weeks to expand hESC in bulk culture depending on the initial passage number of the cells and the number of flasks of cells required. Two days are needed to set up a differentiation experiment. The day before beginning a spin EB experiment, it is necessary to passage the cultures and prepare the plates and medium for the experiment. The duration of differentiation is variable as specified/required by the researcher—up to 30 days.

Acknowledgements This work was supported by the Australian Stem Cell Centre (ASCC), the Juvenile Diabetes Research Foundation (JDRF), and the National Health and Medical Research Council (NHMRC) of Australia.

Literature Cited Conner, D.A. 2000. Mouse embryo fibroblast (MEF) feeder cell preparation. Curr. Protoc. Mol. Biol. 51:23.22.21-23.22.27.

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Costa, M., Sourris, K., Hatzistavrou, T., Elefanty, A.G., and Stanley, E.G. 2007. Expansion of human embryonic stem cells in vitro. Curr. Protoc. Stem Cell Biol. 1:1C.1.1-1C.1.7. Davis, R.P., Ng, E.S., Costa, M., Mossman, A.K., Sourris, K., Elefanty, A.G., and Stanley, E.G. 2007. Targeting a GFP reporter gene to the Mixl1

locus of human embryonic stem cells identitifies primitive streak-like cells and enables isolation of primitive hematopoietic precursors. Blood November 21. Epub. ahead of print. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Hirst, C.E., Ng, E.S., Azzola, L., Voss, A.K., Thomas, T., Stanley, E.G., and Elefanty, A.G. 2006. Transcriptional profiling of mouse and human ES cells identifies SLAIN1, a novel stem cell gene. Dev. Biol. 293:90-103. Keller, G., Kennedy, M., Papayannopoulou, T., and Wiles, M.V. 1993. Hematopoietic commitment during embryonic stem cell differentiation in culture. Mol. Cell Biol. 13:473-486. Kennedy, M., D’Souza, S.L., Lynch-Kattman, M., Schwantz, S., and Keller, G. 2007. Development of the hemangioblast defines the onset of hematopoiesis in human ES cell differentiation cultures. Blood 109:2679-2687. Lacaud, G., Gore, L., Kennedy, M., Kouskoff, V., Kingsley, P., Hogan, C., Carlsson, L., Speck, N., Palis, J., and Keller, G. 2002. Runx1 is essential for hematopoietic commitment at the hemangioblast stage of development in vitro. Blood 100:458-466. Martin, G.R. 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. 78:76347638. Ng, E.S., Azzola, L., Sourris, K., Robb, L., Stanley, E.G., and Elefanty, A.G. 2005a. The primitive streak gene Mixl1 is required for efficient haematopoiesis and BMP4-induced ventral mesoderm patterning in differentiating ES cells. Development 132:873-884. Ng, E.S., Davis, R.P., Azzola, L., Stanley, E.G., and Elefanty, A.G. 2005b. Forced aggregation of defined numbers of human embryonic stem cells into embryoid bodies fosters robust, reproducible hematopoietic differentiation. Blood 106:1601-1603. Current Protocols in Stem Cell Biology

Pick, M., Azzola, L., Mossman, A., Stanley, E.G., and Elefanty, A.G. 2007. Differentiation of human embryonic stem cells in serum free medium reveals distinct roles for bone morphogenetic protein 4, vascular endothelial growth factor, stem cell factor, and fibroblast growth factor 2 in hematopoiesis. Stem Cells. 25:2206-2214. Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399404. Robertson, S.M., Kennedy, M., Shannon, J.M., and Keller, G. 2000. A transitional stage in the commitment of mesoderm to hematopoiesis requiring the transcription factor SCL/tal-1. Development 127:2447-2459. Smith, J., Wardle, F., Loose, M., Stanley, E.G., and Patient, R. 2007. Germ layer induction in ESC— Following the vertebrate road map. Curr. Protoc. Stem Cell Biol. 1:1D.1.1-1D.1.22. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

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Differentiation of Human Embryonic Stem Cells in Adherent and in Chemically Defined Culture Conditions

UNIT 1D.4

Ludovic Vallier1 and Roger Pedersen1 1

Department of Surgery and Cambridge Institute for Medical Research, Addenbrooke’s Hospital, University of Cambridge, Cambridge, United Kingdom.

ABSTRACT Generating fully functional differentiated cells from human embryonic stem cells and achieving this goal using clinically compatible conditions remain major challenges for the stem cell field. The presence of undefined components in standard culture media and protocols (including animal-derived serum, feeder cells, and extracellular matrices) has significantly impeded the achievement of these objectives. Here, we describe culture conditions to differentiate pluripotent cells in adherent conditions and in the absence of stroma cells, feeder cells, conditioned medium, serum, or complex matrices. Importantly, these defined culture conditions are devoid of animal products, thereby eliminating factors that could obscure analysis of developmental mechanisms or render the resulting tissues incompatible with future clinical applications. Curr. Protoc. Stem Cell Biol. 4:1D.4.1C 2008 by John Wiley & Sons, Inc. 1D.4.7.
Keywords: embryonic stem cells r differentiation r pluripotency r chemically defined

The generation of fully functional differentiated cells from human embryonic stem cells (hESCs) and achieving this goal using clinically compatible conditions remain major challenges. The presence of undefined components in standard culture media and protocols (including animal-derived serum, feeder cells, and extracellular matrices) has significantly impeded the achievement of these objectives. This unit describes culture conditions to differentiate pluripotent cells in adherent conditions and in the absence of stroma cells, feeder cells, conditioned medium, serum, or complex matrices. Importantly, these defined culture conditions are devoid of animal products, thereby eliminating factors that could obscure analysis of developmental mechanisms or render the resulting tissues incompatible with future clinical applications. The first step of this approach consists in transferring hESCs (grown initially on feeder cells or in serum-containing media) into a chemically defined medium. The second step consists of adding growth factors or inhibitors of specific signaling pathways to induce the differentiation of pluripotent cells into derivatives of the three primary germ layers: ectoderm, mesoderm, and endoderm.

BASIC PROTOCOL

NOTE: All procedures should be performed under sterile conditions. All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly.

Materials Cultures of hESC cells (grown on feeder or in feeder-free conditions, confluent) Calcium- and magnesium-free phosphate-buffered saline (CMF-PBS; Invitrogen, cat. no. 14190-094) 10 mg/ml human fibronectin (Chemicon, cat. no. FC010)

Current Protocols in Stem Cell Biology 1D.4.1-1D.4.7 Published online March 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01d04s4 C 2008 John Wiley & Sons, Inc. Copyright


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Chemically defined medium (CDM; see recipe) Activin (R&D Systems) FGF2 (R&D Systems) Collagenase solution (see recipe) Growth factors (see recipe) 6- or 12-well plastic plate, tissue culture treated (Corning) 5-ml pipet 15-ml conical centrifuge tube Additional reagents and equipment for immunofluorescence, FACS, and PCR analyses (UNIT 1B.3) Prepare fibronectin-coated plate 1. Add 1.5 ml CMF-PBS to each well of a 6-well plate. 2. Add 15 µl of 10 mg/ml human fibronectin per well of a 6-well plate. For a 12-well plate add 0.5 ml CMF-PBS per well and then 5 µl of fibronectin.

3. Incubate at 4◦ C overnight or 20 min to 1 hr at 37◦ C. 4. Wash each well once with 2 ml CMF-PBS. 5. Add 2 ml of CDM containing 10 ng/ml activin A and 12 ng/ml FGF to each well of a 6-well plate Use 1 ml/well for a 12-well plate. Activin A and FGF2 are used in the medium to maintain the pluripotent status of the hESCs.

Transfer hESCs into feeder-free and chemically defined culture conditions 6. Wait until hESCs grown on feeders reach maximum confluency without differentiation and then start the transfer into chemically defined conditions. To our knowledge, hESCs grown on feeders cannot be used for clinical application even after being transferred in CDM. Only hESCs established on human feeders in a good manufacturing practice (GMP) environment could qualify for clinical use [as long as the BSA contained in the CDM is replaced by human serum albumin (hSA)]. So far only one publication has described hESC lines derived in such conditions as fully compatible with clinical applications (Crook et al., 2007).

7. Wash confluent hESCs once with 2 ml CMF-PBS per well of 6-well plate. 8. Add 3 ml of collagenase solution. 9. Incubate 15 min at 37◦ C. 10. Scrape the hESC colonies off the plate using a 5-ml pipet. 11. Transfer the resulting clumps into a 15-ml tube containing 3 ml of CDM (without growth factors). 12. Gently pipet the resulting clumps up and down (triturate) to dissociate them further. The final size of the clumps is important. The cells will differentiate if the clumps are too small (<200 cells) or too large (>1000 cells).

13. Pellet the clumps by centrifuging 3 min at 200 × g, room temperature. Differentiation of hESC in Adherent and Chemically Defined Conditions

14. Discard the supernatant.

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15. Resuspend the cells in CDM containing 10 ng/ml activin and 12 ng/ml FGF. One confluent well can be divided into 12 wells of the same plate size. In this case resuspend the cells in 12 ml of CDM containing activin and FGF.

16. Plate the hESC clumps at low density (100 to 300 clumps/well) on a fibronectincoated 6-well plate in 2 ml of CDM containing 10 ng/ml activin A and 12 ng/ml FGF2. The total volume for one well of a 6-well plate should be 3 ml.

17. Replace medium the following day with 1 ml per well of fresh CDM containing 10 ng/ml activin A and 12 ng/ml FGF2. One milliliter of medium is sufficient to grow cells in a 6-well plate and, therefore, these plates are more cost effective.

18. Wait 48 hr before adding growth factors or inducing differentiation in these cultures.

Differentiate cultures into ectoderm, mesoderm, or endoderm 19. Add 1 ml CDM, containing growth factors for differentiation, per well of 6- well plate. 10 ng/ml BMP4 10 is known to drive differentiation of hESCs into trophectoderm (Xu et al., 2002). High doses of activin A (100 ng/ml) in combination with Wnt3A (25 ng/ml) in the presence of low levels of fetal bovine serum (0.5%) induce differentiation of hESCs into mesendoderm and definitive endoderm (D’Amour et al., 2005, 2006). FBS is used to induce differentiation of hESCs into mesoendoderm. To our knowledge, there is no alternative published method to drive differentiation of hESCs into this particular germ layer. However, all the other media used in this manuscript are depleted of FBS. Inhibition (see Reagents and Solutions section) of BMP4 signaling using noggin (100 ng/ml; Zhang et al., 2001) and activin A/nodal signaling using Lefty or SB-5431542 (Inman et al., 2002; Smith et al., 2008) has been shown to drive differentiation of hESCs into neuroectoderm progenitors

20. Change the CDM supplemented with growth factors every day for 7 days to obtain a maximum differentiation. 21. Analyze the cultures for expression of the expected markers by immunofluorescence (UNIT 1B.3), FACS (UNIT 1B.3), and PCR analyses (UNIT 1B.3). See Table 1D.4.1 for markers. Table 1D.4.1 Markers for Differentiation of hESC

Cell Type

Inducer

Markers

Pluripotent

Activin A, FGF2

Sox2, Oct-4, nanog

Mesendodem

High-dose activin, Wnt3A, low% FBS

Brachyury, PDGFαR, Mixl1, Eomes

Mesoderm

Brachyury, PDGFαR

Endoderm, primitive

BMP4

Sox7, GATA4, GATA6

Endoderm, definitive

High-dose activin, Wnt3A, low% FBS

Sox17, goosecoid, CXCR4

Neuroectoderm

Noggin, Lefty/SB5431542

Sox2, Sox1, Pax6, N-Cam

Trophectoderm

BMP4

CDX2, hand1, αhCG

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Chemically defined medium (CDM), 500 ml 250 ml Iscove’s modified Dulbecco’s medium (IMDM, Invitrogen; 50% final) 250 ml F-12 medium (Invitrogen; 50% final) 2.5 g bovine serum albumin (BSA, Equibiotech, Europa Bioproducts; 5 mg/ml) or 0.5 g human serum albumin (hSA, Sigma; 1 mg/ml final) 5 ml chemically defined lipids (Invitrogen; cat. no. 11905-031; 1% v/v final) 20 µl monothioglycerol (Sigma; 450 µM final) 350 µl 10 mg/ml insulin (Roche; 7 mg/ml final) 250 µl 30 mg/ml transferrin (Roche; 15 mg/ml final) Store up to 2 weeks at 4◦ C BSA is an essential component of the chemically defined medium (CDM), and its animal origin might represent a drawback for clinical use. However, BSA can be replaced by human serum albumin, which is compatible with clinical applications but which is also less cost effective. Therefore, BSA remains the best solution for large-scale experiments and basic studies.

Collagenase solution, 500 ml 500 ml Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen) 0.5 g collagenase IV (Invitrogen) Store up to 2 weeks at 4◦ C Growth factors (1000× stock solutions) 10 ng/µl activin A (R&D Systems) 4 ng/µl FGF-2 (R&D Systems) 10 ng/µl BMP4 (R&D Systems) 25 ng/µl Wnt3A (R&D Systems) For reconstitution of the growth factors, it is recommended that sterile PBS containing at least 0.1% human serum albumin or bovine serum albumin be added to the vial to prepare a stock solution of no less than 10 µg/ml. For BMP4 only, it is recommended that sterile 4 mM HCl containing at least 0.1% human serum albumin or bovine serum albumin be added to the vial to prepare a stock solution of no less than 10 µg/ml. Activin A, FGF2, BMP4 are recombinant human proteins. Wnt3 is a recombinant mouse protein.

Signaling pathways inhibitors (1000× stock solutions) 10 mM SB5431542 (Tocris) in DMSO Store at −20◦ C SB5431542 is an activin/nodal/TGFβ receptor inhibitor. 25 ng/ml noggin (R&D Systems) For reconstitution of noggin, it is recommended that sterile PBS containing at least 0.1% human serum albumin or bovine serum albumin be added to the vial to prepare a stock solution of no less than 10 µg/ml.

Noggin is a BMP inhibitor. Store at −20◦ C. Differentiation of hESC in Adherent and Chemically Defined Conditions

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COMMENTARY Background Information The chemically defined medium described in this method was developed by Johansson and Wiles (1995). This medium was first used to drive differentiation of mouse ESCs into mesoderm derivatives by using the embryoid body (EB) approach. Importantly, hESCs can be grown over prolonged periods in CDM supplemented with activin and FGF (Vallier et al., 2005; Brons et al., 2007) while maintaining their pluripotency and a normal karyotype. Contrary to their murine counterpart, hESCs possess the ability to differentiate into trophectoderm cells (expressing CDX2, Hand1, and αhCG) in the presence of BMP4. These results have been obtained in the presence of serum and on Matrigel (Xu et al., 2002). Similar results can be obtained using the CDM on fibronectin, with the difference that BMP4 under these conditions also drives differentiation of hESCs into primitive endoderm cells expressing Sox7. Importantly, only the combination of several markers allows the distinction between primitive endoderm cells (expressing Sox7, GATA4, GATA6) and definitive endoderm (expressing Sox17, GSC, CXCR4) from which adult organs including liver and pancreas are derived (Tada et al., 2005; Yasunaga et al., 2005). A number of publications have described the generation of neuroectoderm cells from hESCs (Zhang et al., 2001; Li et al., 2005; Joannides et al., 2007). The culture media described in these publications generally use FGF2 and the absence of any other growth factors including serum to drive differentiation of hESCs toward neuroectoderm progenitors (expressing Sox2, Sox1, and Pax6). Growing hESCs in CDM in the presence of FGF2 and in the absence of activin signaling also results in the generation of neuroectoderm progenitors expressing Sox1 and Pax6. The activin/nodal signaling pathway has been shown to be the essential inducer of mesendoderm differentiation in a large number of species. However, the activin/nodal signaling pathway is also necessary to maintain the pluripotent status of hESCs (Vallier et al., 2004; 2005). The effect of activin/nodal signaling on pluripotency can be redirected to drive differentiation toward endoderm using a high dose of activin (>100 ng/ml) in combination with Wnt3a and low levels of fetal bovine serum (D’Amour et al., 2005, 2006).

Critical Parameters The quality of hESCs is critical. Differentiation background can interfere with the effect of each individual growth factor indicated above. Therefore, it is critically important to start with homogenous populations of undifferentiated hESCs. The size of hESC colonies and culture density affect differentiation. When plated on fibronectin, the hESC clumps should contain at least 200 cells. Smaller colonies may start to differentiate independently of the growth factors added, and larger colonies may have problems attaching. High density of colonies can also influence differentiation and its efficiency. Indeed, hESCs themselves express growth factors (including activin and nodal) that can slow the differentiation or interfere with the effect of the added growth factors. Which hESC lines are used affects differentiation. The efficacy of each method for driving hESC differentiation into a homogeneous population of one particular cell type can vary between different hESC lines. Indeed, there is a growing number of reports that individual hESC lines show different potencies for differentiation for each germ layer. For example, it has been reported that some lines can differentiate more efficiently into endoderm progenitors than others (D’Amour et al., 2005, 2006). BSA is an essential component of the CDM but its animal origin might represent a major limitation for clinical application. To avoid this drawback, BSA can be replaced by human serum albumin (hSA) or by the chemical compound polyvinyl alcohol.

Troubleshooting hESCs can have some difficulty attaching on fibronectin-coated plates. Adhesion of hESCs to fibronectin can be improved by waiting 48 hr after passaging before adding fresh CDM.

Anticipated Results Extraembryonic tissues Differentiated cells will progressively appear after 4 days of BMP4 treatment, and pluripotent cells expressing Oct-4 will totally disappear after 7 days. Extraembryonic differentiation can be monitored by the expression

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of the primitive endoderm marker, Sox7, and the trophectoderm marker, CDX2. Neuroectoderm The absence of activin and BMP signaling induces differentiation of hESCs toward the neuroectoderm lineage in 4 or 5 days. However, expression of the neuroectoderm progenitor markers, Pax6 and Sox1, will only appear after 5 to 6 days of treatment. Importantly, Sox2, expressed in combination with Oct-4 and Nanog, marks pluripotent cells. However, Sox2 expression remains during early stages of neuroectoderm differentiation while Oct-4 and Nanog expression disappears. Homogeneity of differentiation can be validated by FACS analysis for the expression of the pan neuronal marker, NCam. Mesendoderm Mesoderm and endoderm differentiation is marked by the successive expression of markers. Brachyury expression first indicates the differentiation of hESCs into mesendoderm. Then, expression of Sox17, Goosecoid and CXCR4 appears during the commitment of these mesendodem progenitors to definitive endoderm. Brachyury expression remains only in mesoderm cells. Homogeneity of differentiation can be validated by FACS analysis for the expression of the definitive endoderm marker, CXCR4, and the mesendoderm/mesoderm marker, PDGFαR. A differentiation is considered as homogenous when 70% of the cells generated express a particular marker (i.e., CXCR4 for endoderm, PDGFαR for mesendoderm, and N-CAM for neuroectoderm).

Time Considerations A fully differentiated population (i,e., absence of expression of pluripotency markers) is usually obtained after at least 7 days of treatment. However, homogeneity of differentiation could vary depending on the human ES cell line used and the time of treatment can be extended to improve the differentiation.

Literature Cited Brons, I.G., Smithers, L.E., Trotter, M.W., RuggGunn, P., Sun, B., Chuva de Sousa Lopes, S.M., Howlett, S.K., Clarkson, A., Ahrlund-Richter, L., Pedersen, R.A., and Vallier, L. 2007. Derivation of pluripotent epiblast stem cells from mammalian embryos. Nature 448:191-195. Differentiation of hESC in Adherent and Chemically Defined Conditions

Crook, J.M., Peura, T.T., Kravets, L., Bosman, A.G., Buzzard, J.J., Horne, R., Hentze, H., Dunn, N.R., Zweigerdt, R., Chua, F., Upshall, A., and Colman, A. 2007. The generation of six

clinical-grade human embryonic stem cell lines. Cell Stem Cell 1:490-494. D’Amour, K.A., Agulnick, A.D., Eliazer, S., Kelly, O.G., Kroon, E., and Baetge, E.E. 2005. Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23:1534-1541. D’Amour, K.A., Bang, A.G., Eliazer, S., Kelly, O.G., Agulnick, A.D., Smart, N.G., Moorman, M.A., Kroon, E., Carpenter, M.K., and Baetge, E.E. 2006. Production of pancreatic hormoneexpressing endocrine cells from human embryonic stem cells. Nat. Biotechnol. 24:13921401. Inman, G.J., Nicolas, F.J., Callahan, J.F., Harling, J.D., Gaster, L.M., Reith, A.D., Laping, N.J., and Hill, C.S. 2002. SB-431542 is a potent and specific inhibitor of transforming growth factorbeta superfamily type I activin receptor-like kinase (ALK) receptors ALK4, ALK5, and ALK7. Mol. Pharmacol. 62:65-74. Joannides, A.J., Fiore-Heriche, C., Battersby, A.A., Athauda-Arachchi, P., Bouhon, I.A., Williams, L., Westmore, K., Kemp, P.J., Compston, A., Allen, N.D., and Chandran, S. 2007. A scaleable and defined system for generating neural stem cells from human embryonic stem cells. Stem Cells 25:731-737. Johansson, B.M. and Wiles, M.V. 1995. Evidence for involvement of activin A and bone morphogenetic protein 4 in mammalian mesoderm and hematopoietic development. Mol. Cell Biol. 15:141-151. Li, X.J., Du, Z.W., Zarnowska, E.D., Pankratz, M., Hansen, L.O., Pearce, R.A., and Zhang, S.C. 2005. Specification of motoneurons from human embryonic stem cells. Nat. Biotechnol. 23:215221. Smith, J.R., Vallier, L., Lupo, G., Alexander, M., Harris, B., and Pedersen, R.A. 2008. Inhibition of Activin/Nodal signaling promotes differentiation of human embryonic stem cells into neuroectoderm. Dev. Biol. 313:107-117. Tada, S., Era, T., Furusawa, C., Sakurai, H., Nishikawa, S., Kinoshita, M., Nakao, K., and Chiba, T. 2005. Characterization of mesendoderm: A diverging point of the definitive endoderm and mesoderm in embryonic stem cell differentiation culture. Development 132:43634374. Vallier, L., Rugg-Gunn, P.J., Bouhon, I.A., Andersson, F.K., Sadler, A.J., and Pedersen, R.A. 2004. Enhancing and diminishing gene function in human embryonic stem cells. Stem Cells 22:2-11. Vallier, L., Alexander, M., and Pedersen, R.A. 2005. Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell Sci. 118:4495-4509. Xu, R.H., Chen, X., Li, D.S., Li, R., Addicks, G.C., Glennon, C., Zwaka, T.P., and Thomson, J.A. 2002. BMP4 initiates human embryonic stem cell differentiation to trophoblast. Nat. Biotechnol. 20:1261-1264.

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Yasunaga, M., Tada, S., Torikai-Nishikawa, S., Nakano, Y., Okada, M., Jakt, L.M., Nishikawa, S., Chiba, T., and Era, T. 2005. Induction and monitoring of definitive and visceral endoderm differentiation of mouse ES cells. Nat. Biotechnol. 23:1542-1550. Zhang, S.C., Wernig, M., Duncan, I.D., Brustle, O., and Thomson, J.A. 2001. In vitro differentiation of transplantable neural precursors from human embryonic stem cells. Nat. Biotechnol. 19:O1129-1133.

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Isolation and Differentiation of Xenopus Animal Cap Cells

UNIT 1D.5

Takashi Ariizumi,1 Shuji Takahashi,1 Te-chuan Chan,2 Yuzuru Ito,3 Tatsuo Michiue,1 and Makoto Asashima1, 2, 3 1

University of Tokyo, Tokyo, Japan Japan Science and Technology Agency, Tokyo, Japan 3 Organ Development Research Laboratory, National Institute of Advanced Industrial Science and Technology, Ibaraki, Japan 2

ABSTRACT Xenopus is used as a model animal for investigating the inductive events and organogenesis that occur during early vertebrate development. Given that they are easy to obtain in high numbers and are relatively large in size, Xenopus embryos are excellent specimens for performing manipulations such as microinjection and microsurgery. The animal cap, which is the area around the animal pole of the blastula, is destined to form the ectoderm during normal development. However, these cells retain pluripotentiality and upon exposure to specific inducers, the animal cap can differentiate into neural, mesodermal, and endodermal tissues. In this sense, the cells of the animal cap are equivalent to mammalian embryonic stem cells. In this unit, the isolation and differentiation of animal cap cells, the so-called animal cap assay, is described. Useful methods for analyzing the mechanism of animal cap differentiation at the molecular level are also described. Curr. Protoc. Stem C 2009 by John Wiley & Sons, Inc. Cell Biol. 9:1D.5.1-1D.5.31.  Keywords: animal cap r pluripotency r activin r retinoic acid r induction r organogenesis r Xenopus laevis

INTRODUCTION Xenopus laevis, an anuran amphibian, has many advantageous features as an animal model over other vertebrates: (1) fertilized eggs are easily obtained by hormonestimulated mating or in vitro fertilization; (2) the developmental rate of these eggs can be regulated thermally; (3) the embryos are large enough to allow surgical manipulations; and (4) isolated embryonic tissues can be easily cultured in a simple salt solution, such as Steinberg’s solution. Therefore, the Xenopus embryo has been used as a resource for understanding the mechanism of early vertebrate development. In blastula-stage embryos, a circular area with the pigmented or animal pole at its center is called the animal cap. This region is fated to become the ectoderm during normal development; its dorsal side forms neural tissues and its ventral side becomes epidermis. The animal cap remains spherical and forms an irregular-shaped epidermis, which is referred to as atypical epidermis, when cultured in isolation. However, the animal cap is competent to respond to inducing molecules, whereby it can form neural, mesodermal, and endodermal tissues. In this sense, the cells of the animal cap are equivalent to mammalian embryonic stem cells. Based on this pluripotency of animal cap cells, a simple and reliable in vitro assay system, termed the animal cap assay, has been devised. In the animal cap assay, investigators can test numerous factors in solution, and can estimate their inducing activities both qualitatively and quantitatively. Moreover, the synergistic effect of two or more factors can be examined by combining them in the

Current Protocols in Stem Cell Biology 1D.5.1-1D.5.31 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01d05s9 C 2009 John Wiley & Sons, Inc. Copyright 

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solution. The competencies of reacting tissues can be analyzed as a model system, in which animal caps of different age or size are treated with various concentrations of an inducer for a defined time period. By combining microinjection techniques and the animal cap assay, it is also possible to assess the activities of the injected genes or their RNAs. Furthermore, the animal caps removed from embryos injected with a cell-lineage tracer at the early cleavage stages can serve as donors in transplantation experiments. In this unit, a main protocol for the animal cap assay (see Basic Protocol 1) is described, and protocols with possible modifications (see Alternate Protocols 1 and 2) are also provided. Before performing the animal cap assay, investigators must obtain fertilized eggs and embryos (see Support Protocols 1 and 2) and prepare special instrumentation that is required for micromanipulations (see Support Protocol 3). Many specific antibodies are available for the identification of the induced tissues in the animal cap explants (see Support Protocol 6). In addition, many practical methods using molecular biological techniques, such as RT-PCR (see Support Protocol 5) and whole-mount in situ hybridization (see Support Protocol 7), have been established for Xenopus embryos. Techniques are described to facilitate analyses of the inductive events for animal caps (see Support Protocols 4 through 7). Investigators are expected to select and combine these protocols according to the design and purpose of individual experiments. BASIC PROTOCOL 1

Isolation and Differentiation of Xenopus Animal Cap Cells

ANIMAL CAP ASSAY The outline of the animal cap assay is shown in Figure 1D.5.1. In this protocol, the membrane-free blastula is placed with the animal pole facing upwards. The animal cap area is squarely dissected using a fine tungsten needle. The test solutions of soluble inducers (e.g., activin and fibroblast growth factor) are tested for their inducing activities by adding them to the animal caps in a saline solution, such as Steinberg’s solution. The procedure for dissecting the animal cap from the blastula is shown in Video 1.

Figure 1D.5.1 Outline of the animal cap assay. An animal cap removed from a blastula is immersed in a saline solution that contains various concentrations of inducer. In the absence of inducer, the cap forms a cluster of epidermis, termed atypical epidermis. The differentiation of mesodermal tissues, such as the notochord and muscle, indicates the mesoderm-inducing activity of the inducer, whereas the differentiation of neural tissues, such as the brain and eyes, indicates the neural-inducing activity of the inducer.

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Materials Blastula embryos at developmental stages 8 or 9 (Fig. 1D.5.2) Steinberg’s solution (SS; see recipe) 0.1% (w/v) bovine serum albumin in SS (pH 7.4; 0.1% BSA-SS) Test solutions (e.g., such as activin and fibroblast growth factor dissolved in 0.1% BSA-SS) Operating dishes, transfer pipets, and tungsten needles (see Support Protocol 3) Low-adhesion, 24-well tissue culture plate (Sumitomo Bakelite, cat. no. MS-80240) 20◦ to 22◦ C incubator 1. After removing the vitelline membrane, place the embryos with the animal pole side up in an operating dish filled with SS. 2. Trim both sides of the embryo with the tungsten needle. 3. Insert the needle into the blastocoel from one side, and divide the vegetal hemisphere (endoderm) by pushing down the needle. 4. Reverse the sheet of the animal cap having endodermal cell masses at each end, and dissect them from the sheet. 5. Trim the animal cap carefully to an area of 0.5 × 0.5 mm, to eliminate adjacent marginal zone cells. 6. Transfer the cap to the test solution, and place it so that the inner blastocoel side is oriented towards the top. Test solutions are prepared in a low-adhesion, 24-well tissue culture plate. BSA is added to the solutions to a final concentration of 0.1% (w/v), to avoid adsorption of inducer(s) to the plastic surfaces.

Figure 1D.5.2 Temperature-dependent early development of Xenopus embryos. Within the normal tolerance range (18◦ to 24◦ C), it is possible to retard or accelerate the rate of embryonic development without altering the developmental processes. Embryonic and Extraembryonic Stem Cells

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7. After exposure to the test solutions for a defined time period, wash the animal caps in 5 ml SS with gentle pipetting and culture them in 1 ml fresh SS at 20◦ to 22◦ C. One may expose the explants in the test solution for the entire culture period (2 to 3 days). Troubleshooting strategies, examples of Anticipated Results, and Time Considerations related to the animal cap assay are described in the Commentary. SUPPORT PROTOCOL 1

OBTAINING FERTILIZED EGGS AND MEMBRANE REMOVAL Xenopus laevis can be induced to mate naturally at 3-month intervals by the injection of human chorionic gonadotropin (hCG). A fully mature female lays several thousand eggs at one spawning. The embryos are surrounded by a jelly coat and vitelline membrane. These membranes must be removed before any manipulation of the embryos can occur. Jelly coats are usually dissolved chemically, whereas vitelline membranes are manually removed with two pairs of forceps.

Materials hCG dissolved in saline (0.9% NaCl) at a concentration of 2000 U/ml Fully mature male and female frogs (Xenopus laevis or X. borealis) Steinberg’s solution (SS; see recipe) Dejelling solution (CSS): 4.5% (w/v) cysteine-HCl in SS (pH 7.8), prepare fresh Sterilized 1-ml syringe with 26-G needle 10- to 15-liter container Thin plastic card Large-bore pipet (∼5-mm diameter) Sterilized beakers (100-ml) Operating dishes, transfer pipets, and two pairs of watchmaker’s forceps (see Support Protocol 3) Mate frogs naturally and collect eggs 1. Load 2000 U/ml hCG into a sterile 1-ml syringe with a 26-G needle attached. Insert the needle into each frog beneath the skin of the thigh and push it forward beyond the “stitch” marks (see Fig. 1D.5.3A). 2. When the tip of needle has reached the dorsal lymph sac, inject the animal (one male and one female frog) with 600 U (0.3 ml) of hCG. For more reliable results, it is advisable to inject the male with half (300 U) of the hCG dosage at least 6 hr before the final injection for mating. Penetration of the dorsal lymph sac is easily recognized from the outside because the skin is thin and very loose.

3. Place the frogs together in a 10- to 15-liter container that is filled with dechlorinated water to a depth of ∼10 cm, and incubate overnight at 20◦ to 22◦ C. The fertilized eggs can be obtained ∼12 hr after the injection.

4. Using a thin plastic card, scrape off the fertilized eggs that adhere to the bottom of the container, and collect them using the large-bore pipet (see Fig. 1D.5.3B).

Isolation and Differentiation of Xenopus Animal Cap Cells

The early stages of development are influenced by environmental conditions, especially the water temperature. The temperature tolerance of Xenopus embryos is given in Figure 1D.5.2. Within the normal tolerance range, it is possible to retard or accelerate the developmental rate without altering the developmental processes. The table that contains the normal range of values (Nieuwkoop and Faber, 1967) is available for the staging of embryos.

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Figure 1D.5.3 Obtaining eggs by hormone-stimulated natural mating. (A) Fertilized eggs are obtained by the injection of hCG into the dorsal lymph sacs of the male and female. (1) A 1-ml syringe filled with hCG; (2) “stitch” marks (indicated by a white dotted line); (3) the dorsal lymph sac; and (4) the cloaca. (B) The laying of fertilized eggs begins at the bottom of the container ∼12 hr after hCG injection. (1) Male; (2) female; (3) a large-bore pipet; and (4) a thin plastic card for egg collection.

Remove jelly coat 5. Collect the embryos in a sterilized 100-ml beaker and wash them with 50 ml SS. 6. Discard the SS and add 50 ml CSS. 7. Remove jelly coats by gently swirling for a few minutes (see Video 2). The jelly coats fall off and the embryos begin to pack closely together. Since prolonged exposure to CSS will damage the embryos, the dejellied embryos must be washed immediately in SS.

8. Decant the CSS and immediately rinse at least ten times in 50 ml SS with gentle swirling.

Remove vitelline membrane 9. Select embryos according to the developmental table, and place them into an operating dish that contains 50 ml SS. 10. For the animal cap assay, hold the blastula embryo upside down, and then quickly grasp and tear the membrane using two pairs of watchmaker’s forceps (see Video 1). It is not a problem if a few vegetal cells are injured when the membrane is grasped.

11. Place the membrane-free blastula with the animal pole facing upwards. 12. Dissect the animal cap area using a fine tungsten needle (see Basic Protocol 1).

IN VITRO FERTILIZATION AND RAPID REMOVAL OF THE JELLY COAT In vitro fertilization is advantageous, particularly in the microinjection study, for synchronizing embryos to the same developmental stage. This protocol is concerned with in vitro fertilization and rapid removal of the jelly coat from the fertilized egg. Injection occurs within 30 min, causing cleavage to begin within 30 min and continues every 30 min, without intervals, making these techniques suitable for microinjection studies.

SUPPORT PROTOCOL 2

Materials Fully mature male and hCG-primed female frogs (Xenopus laevis) Anesthetic: 0.1% (w/v) ethyl 3-aminobenzoate methanesulfonate salt (Tricaine/MS222; Sigma) in tap water (not distilled water) DeBoer’s solution (DB; see recipe) FBS

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Dejelling solution: 1% (w/v) sodium thioglycollate in SS (pH 6.0) 1 M NaOH Steinberg’s solution (SS; see recipe) Surgical board Forceps Scissors 60-mm dishes 15-ml conical tubes Pasteur pipets with tipfused by a flame Isolate testes 1. Immerse frog in 1 liter of anesthetic and allow 20 to 30 min for the anesthetic to take effect. hCG-priming can be performed on other Xenopus such as X. tropicalis and X. borealis, but the injection volume, timing, and the number of times are different. Therefore, these protocol parameters apply only to Xenopus laevis.The size of the frog is not of concern. Instead of anesthetic, ice water can be used. Do not leave the frogs in the anesthetic for longer than is necessary for anesthesia. If only one testis is to be used within 2 weeks, remove one testis, stitch up wound, and revive frog to use in a later experiment.

2. Place the anesthetized frog belly up on a surgical board. Pick up the belly skin using a pair of forceps and cut the skin open with scissors. Then, cut the abdominal muscles. Be careful not to cut the large blood vessel running along the midline.

3. Pull out the fat body. Remove the testes and the fat body from the kidney and place in a 60-mm dish, and then remove the fat body from the testes. The white testis is located at the boundary between the fat body and the kidney.

4. Wash the testes in 10 ml DB and then wipe the blood from the vessels using a paper towel. 5. Place the testes in 10% FBS/90% DB and store at 4◦ C. Testes can be stored for 1 to 2 weeks. Testis-removed frogs are sacrificed and stored at −20◦ C.

Prepare sperm suspension 6. Mince with scissors one-half of the testes in a droplet of DB. 7. Suspend the sperm in 5 to 10 ml of DB. Transfer the solution into a 15-ml conical tube and store on ice. This sperm suspension can be used for several hours.

Collect eggs 8. Confirm that the hCG-primed female (see Basic Protocol 1) is laying eggs from the cloaca. 9. Hold the frog gently with both hands (see Fig. 1D.5.4A). Push the region near the cloaca with thumb and forefinger. Collect eggs in a 60-mm dish. Maintain pressure but do not squeeze frog during egg collection. If the frog kicks your hands with its claws, press the head of the frog tightly using the palm and ring finger or little finger. Isolation and Differentiation of Xenopus Animal Cap Cells

Fertilize eggs 10. Add two or three drops of the sperm suspension onto the collected eggs.

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Figure 1D.5.4 Obtaining fertilized eggs by in vitro fertilization. (A) Confirm that hCG-primed female is laying eggs from the cloaca. Hold the frog gently with both hands and push the region near the cloaca with the thumb and forefinger. Eggs are collected in a 60-mm dish. (B) After adding two or three drops of the sperm suspension and a few drops of DB to the collected eggs, mix and spread them into a single layer on the dish using a Pasteur pipet with flame-fused tip.

11. Mix thoroughly and gently using a Pasteur pipet with tip fused by a flame (see Fig. 1D.5.4B). If this proves difficult due to the viscosity of the jelly, add a few drops of DB and spread using the Pasteur pipet with tip fused by a flame (see Fig 1D.5.7C) into a single layer on the dish.

12. After the eggs have been in contact with sperm for 2 min, pour 10 ml distilled water over the eggs. When the salt concentration is reduced by dilution, sperm start to move into the jelly and towards the eggs. Minutes later, contraction of the animal hemisphere (the pigmented region) of the egg, which is the first sign of fertilization, should occur. The first cleavage occurs 90 min later at 23◦ C.

Remove jelly coat rapidly 13. Discard the water from the dish that contains the fertilized eggs. Pour 10 ml of dejelling solution into the dish. 14. Add 300 to 500 μl of 1 M NaOH to increase the pH to 10 to 10.5. Shake and rotate vigorously as soon as possible. Decant this solution when the jelly coat is dissolved (this takes ∼30 sec). Do not discard all of the solution. Hold the dish at an angle, and add 10 ml SS from the opposite side. 15. Shake and rotate the dish again for 15 sec, and then discard the solution by decanting, and add 10 ml SS. 16. Wash the fertilized eggs three to four times with 10 ml SS until the pH of the solution reaches 7.4. 17. Culture the dejellied eggs in 10 ml SS in a 60-mm dish at 20◦ to 22◦ C.

PREPARATION OF MICROMANIPULATION TOOLS The equipment required for microsurgery is illustrated in Figure 1D.5.5. Operations are usually performed on a clean bench. A binocular microscope with 10× oculars and 1× to 4× objectives, and an illuminator (fiber-optic light is preferable) are needed. The preparation and sterilization of the manipulation tools are described here.

Forceps and tungsten needles Two pairs of watchmaker’s forceps (e.g., Fontax no. 5) are required to remove the vitelline membrane, which lies close to the embryos. The forceps are heat-sterilized for

SUPPORT PROTOCOL 3

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Figure 1D.5.5 Instruments for removing and handling the animal caps. The instruments needed for the animal cap assay are: (1) clean bench; (2) binocular microscope; (3) fiber-optic light; (4) Steinberg’s solution; (5) operating dish; (6) small dishes; (7) samples; (8) tissue culture plate; (9) watchmaker’s forceps for removing the vitelline membrane; (10) tungsten needles for dissecting animal cap tissues; (11) transfer pipets for handling embryos and animal caps.

2 hr at 180◦ C. For the dissection of embryonic tissues, electrolytically sharpened tungsten needles are used. They are durable, can be resharpened, and can be heat-sterilized. 1. Cut 0.2-mm tungsten wire into a 2-cm-length piece using pliers. 2. Mount the wire on a 10-cm × 3-mm soft glass tubing in a flame. 3. Bend the wire at a right angle, at ∼3 to 5 mm from its end. 4. Sharpen the wire end using 5 M NaOH and a dry cell (9V). By placing the negative pole on a carbon point in the NaOH solution and attaching the positive pole to the tungsten wire, repeated dipping into the solution will sharpen the wire to a fine point.

Transfer pipets Pasteur pipets are used for making transfer pipets. 1. Flame the Pasteur pipet at its center and draw it out at a right angle. 2. For transferring embryos, cut pipets with 2-mm diameter using an ampule cutter and smooth the cut edge in a small flame. Similarly, small transfer pipets for animal cap explants are made by cutting the tapered Pasteur pipets a 0.5- to 1-mm diameter. 3. Heat-sterilize 2 hr at 180◦ C, and use together with an ordinary silicon nipple, sterilized in 70% ethanol.

Operating dishes Operations are carried out in 90-mm glass dishes. The base of the dish should be lined with 3% (w/v) agar, to prevent the embryonic tissues from sticking to the glass surface. 1. Dissolve 3 g agar in 100 ml distilled water while heating in a microwave oven, to produce about ten operating dishes. 2. Pour a thin layer (∼10 ml) of molten agar over the base of each dish, and allow it to cool. Isolation and Differentiation of Xenopus Animal Cap Cells

3. Wrap the dishes in aluminum foil, autoclave 20 min at 120◦ C, and allow dishes to harden upon cooling in a horizontal position. These dishes can be stored for a few months at 4◦ C. Store dishes upside down to reduce the formation of condensation on the agar surface.

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Current Protocols in Stem Cell Biology

MULTIPLE TREATMENTS OF ANIMAL CAPS FOR KIDNEY AND PANCREAS INDUCTION

ALTERNATE PROTOCOL 1

Activin induces animal caps to differentiate into muscle, notochord, gut, neural tissues, and other tissues. In combination with another bioactive factor, retinoic acid, activin induces the generation of the pronephros (embryonic kidney) and pancreas. Although retinoic acid does not have inducing activity per se, it modifies the direction of the differentiation of animal caps induced by activin. This protocol describes the treatment of animal caps with activin and retinoic acid.

Materials Late-blastula embryos at developmental stage 9 (Fig. 1D.5.2) 0.1% (w/v) BSA in SS, pH 7.4 (0.1% BSA-SS; see recipe for SS) Retinoic acid stock solution (10−2 M): 3 mg all-trans retinoic acid (Sigma, cat. no. R2625) dissolved in 1 ml DMSO or ethanol Test solution 1: 10 μl retinoic acid stock solution plus 990 μl of 10 ng/ml activin in 0.1% BSA-SS Test solution 2: 100 ng/ml activin in 0.1% BSA-SS Test solution 3: 10 μl retinoic acid plus 990 μl of 0.1% BSA-SS Operating dishes, transfer pipets, and two pairs of watchmaker’s forceps (see Support Protocol 3) Low-adhesion, 24-well tissue culture plate (Sumitomo Bakelite, cat. no. MS-80240) ◦ 20 C incubator Induce pronephros 1. Isolate animal caps (see Basic Protocol 1). 2. Transfer ten caps immediately to 1 ml test solution 1 in a well of a 24-well tissue culture plate. Either DMSO or ethanol can be used to dissolve retinoic acid, although ethanol will reduce the solubility of retinoic acid. The presence of BSA in SS prevents the animal caps from adhering to the surface of the tissue culture plate.

3. Incubate 3 hr at 20◦ C. 4. Wash the caps two times for 5 min in 5 ml of 0.1% BSA-SS. 5. Place ten caps in 1 ml of 0.1% BSA-SS in a well of a 24-well tissue culture plate and culture 3 days at 20◦ C. Be careful to keep the caps apart from each other using forceps or a tungsten needle. Formation of pronephric tubules can be observed inside the thin epidermal vesicle after 4 days of culture. Pronephric differentiation is confirmed by histological examination and expression of specific marker genes.

Perform time-lag treatment for pancreas induction 6. Transfer ten caps immediately to 1 ml of test solution 2 in a well of a 24-well tissue culture plate and incubate 1 hr at 20◦ C. 7. Wash the caps two times for 5 min in 5 ml of 0.1% BSA-SS. 8. Incubate caps in 1 ml of 0.1% BSA-SS for 5 hr at 20◦ C. 9. Transfer caps to 1 ml test solution 3 and incubate 1 hr at 20◦ C. 10. Wash caps two times for 5 min in 1 ml of 0.1% BSA-SS.

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11. Culture in 1 ml of 0.1% BSA-SS for 3 days at 20◦ C. Be careful to keep the caps apart from each other using forceps or tungsten needle. Pancreatic differentiation can be characterized by histological examination and expression of molecular markers such as pdx 1 and insulin. ALTERNATE PROTOCOL 2

DISSOCIATION/REAGGREGATION OF ANIMAL CAPS FOR HEART INDUCTION Animal caps can be dissociated into individual cells by exposure to Ca2+ /Mg2+ -free saline. The cellular adhesion of the caps is loosened within ∼20 min and the cells can be dispersed by gentle pipetting. The dispersed cells are competent for responding to inducers, such as activin, and form reaggregates upon the addition of calcium ions to the test solution or culture medium. This dissociation/reaggregation procedure can be applied to various studies, such as analyses of cell-to-cell interactions, the response of a single cell to an inducing stimulus, and the competencies of the inner and outer cells of the multilayered animal caps. As an example, using the dissociation/reaggregation technique, this protocol describes in vitro heart induction from animal cap explants (see Fig. 1D.5.6).

Materials Mid-blastula embryos at developmental stage 8 (Fig. 1D.5.2) Steinberg’s solution (SS; see recipe) 0.1% (w/v) bovine serum albumin in SS, pH 7.4 (0.1% BSA-SS) 0.1% (w/v) BSA in Ca2+ /Mg2+ -free SS, pH 7.4 (0.1% BSA-CMFSS) Activin solution: 100 ng/ml activin dissolved in 0.1% BSA-SS Operating dishes, transfer pipets, and tungsten needles (see Support Protocol 3) Low-adhesion, 96-well tissue culture plates with concave (U-shaped)-well bottoms (Sumitomo Bakelite, cat. no. MS-30960) 1. Collect five animal caps (0.5 mm × 0.5 mm) from mid-blastula embryos.

Isolation and Differentiation of Xenopus Animal Cap Cells

Figure 1D.5.6 In vitro heart induction using the dissociation/reaggregation protocol. The cellular adhesion of the caps is loosened in CMFSS and the cells can be dispersed by gentle pipetting. The dissociated cells begin to form a reaggregate in SS that contains Ca2+ and 100 ng/ml activin.

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2. Transfer the animal caps to a small operating dish filled with 0.1% BSA-CMFSS, to eliminate Ca2+ and Mg2+ cations transferred from the operating dish. The following steps are performed at 20◦ C in a low-adhesion, 96-well tissue culture plate with U-shaped well bottoms. BSA at 0.1% (w/v) should be added to all solutions to prevent dissociated cells from sticking to the plastic surfaces.

3. Place five caps into a single well that contains 100 μl of 0.1% BSA-CMFSS. 4. Incubate for 20 min at room temperature to disrupt cell adhesion. 5. Replace 0.1% BSA-CMFSS with 100 μl of activin solution and disperse the cells by gentle pipetting. 6. After incubation in the activin solution for 5 hr at room temperature, wash the newly formed spherical “reaggregates” in 5 ml of 0.1% BSA-SS, to eliminate activin. 7. Incubate each reaggregate in a single well filled with 200 μl of 0.1% BSA-SS. The reaggregates will begin to beat rhythmically within 3 days at 20◦ C (see Video 3).

MICROINJECTION OF mRNA FOR ANIMAL CAP ASSAY Animal cap cells are competent to respond to various signaling molecules and transcription factors. Since animal cap cells are formed from fertilized eggs, gene overexpression or downregulation can be achieved by microinjection at an early stage (1-cell or 2-cell stage). In Support Protocol 2, in vitro fertilization and rapid removal of the jelly layer are described. In vitro fertilization and rapid removal of the jelly layer save time in the preparative process for microinjection. The first cleavage requires 90 min and the second and subsequent cleavages take 30 min, without interval, making these methods effective for microinjection.

BASIC PROTOCOL 2

Materials (see Fig. 1D.5.7) Synthetic RNA of interest 5% (w/v) Ficoll in SS In vitro fertilized eggs (see Support Protocol 2) Steinberg’s solution (SS; see recipe) Glass needles Microloader tip (Eppendorf, cat. no. 5242 956.003) Microinjection capillary (e.g., Narishige G-1) Micromanipulator (e.g., Marzhauser MM33) and support base (Drummond Scientific) Microinjector (e.g., PLI-100/-90 Pico-Injector, Harvard/Medical Systems) Microscope Air compressor (e.g., oil-free BEBICON, Hitachi or N2 gas cylinder) 60-mm glass dishes Stainless-steel mesh Pasteur pipets Hair loop or polished forceps 6-well plates, optional Prepare RNA 1. Load the synthetic RNA into a glass needle from behind with a microloader tip, a special fine pipet tip for filling the microinjection capillary. The pCS2+ vector and derivatives thereof are recommended for RNA synthesis (http://sitemaker.umich.edu/dlturner.vectors). These multipurpose expression vectors are very effective for the production of proteins and are used widely in studies on Xenopus

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Figure 1D.5.7 Equipment for microinjection and artificial insemination. (A) Equipment for microinjection: (1) microinjector, (2) binocular microscope and illuminator, (3) manipulator, (4) air compressor, (5) Ficoll solution and Steinberg’s solution, (6) tissue culture plate. (B) A macrophotograph of the end of the glass needle. (C) The instruments needed for microinjection and in vitro fertilization are: (1); Pasteur pipet with flame-fused tip for spreading the fertilized eggs on the dish; (2) transfer pipet for handling embryos; (3) a stainless steel mesh for aligning the embryos; (4) scissors and watchmaker’s forceps.

and zebrafish. The pCS2+ vector contains a strong enhancer/promoter (simian CMV IE94) followed by a polylinker and the SV40 late polyadenlyation site. An SP6 promoter is present in the 5’-untranslated region of the mRNA from the sCMV promoter, and a NotI restriction enzyme site is located after the SV40 late polyadenlyation site, allowing in vitro RNA synthesis of sequences cloned into the polylinker. The mMESSAGE mMACHINE SP6 kit (Ambion) is recommended for the 5’-capped mRNA synthesis. In vitro transcription should be carried out according to the manufacturer’s instructions. For RNA purification, a phenol/chloroform extraction plus double isopropanol precipitation or the RNeasy Mini Kit (Qiagen) for samples for microinjection (see the manufacturer’s instructions for mMESSAGE mMACHINE) is recommended. Synthetic RNA is dissolved in RNase-free water and stored at −20◦ C or −80◦ C. Highly purified RNA can be injected at dosages of up to 2 to 5 ng per embryo. The most effective RNA samples, including those from the Xwnt-8 and Xnr5 genes, are used, and activin can be used at dosages of 100 fg to 10 pg per embryo. The glass needle can be made from a glass capillary (e.g., Narishige G-1) using a glass puller (e.g., Narishige PN-30)

2. Attach the RNA-loaded glass needle to the needle holder connected to the micromanipulator and microinjector. 3. Break off the glass needle tip at a diameter of 5 to 10 μm under the microscope. Inject air and let the air out of the glass needle tip. 4. Inject RNA solution into the air. Adjust the microinjection volume using air pressure and time. Measure the diameter of the sphere using the eyepiece micrometer and calculate the injected volume (v = 4/3πr3 ). A low-volume injection (5 to 10 nl/egg) has no effect on embryo development. In general, the conditions of 35 psi and 0.2 sec produce good results with a needle tip of 5- to 10-μm (depending on the shape of the needle) diameter. Use the balance function to block the capillary phenomenon, and adjust the boundary between the RNA solution in the tip and Ficoll solution in 60-mm dish (see step 5). Isolation and Differentiation of Xenopus Animal Cap Cells

Carry out microinjection 5. Fill a 60-mm glass dish that contains a stainless steel mesh with 5% Ficoll in SS and transfer the eggs using a transfer pipet.

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6. Adjust the orientation of the egg with a hair loop or polished forceps under the microscope. Orient the injection side of the egg towards the microinjection needle tip. 7. Adjust the needle tip to the injection point and prick the blastomere through the vitelline envelope with the needle. Inject RNA solution into the Ficoll solution, while checking the flow (because of refractive index of Ficoll solution, this is easy to check). Occasionally, there are problems with drying of the injection solution or sticking of debris in the needle tip; if this occurs, change needle or re-break needle tip.

8. Inject RNA into the single blastomere of a 1-cell embryo. 9. Withdraw the needle tip and move on to the next egg. 10. After injection, transfer the injected egg to another dish or 6-well plate that contains 5% Ficoll in SS with a transfer pipet. 11. When the injected embryos reach the blastula stage, dissect the animal caps from them (see Basic Protocol 1) and use the animal caps for assays.

HISTOLOGICAL EXAMINATION OF ANIMAL CAP EXPLANTS For interpreting the results of the animal cap assay, it is essential to prepare histological sections of the explants. This process provides accurate information on cell differentiation within the animal caps. Standard protocols, including Bouin’s fluid fixation, paraffin embedding, sectioning, and hematoxylin/eosin staining can be used. The equipment required for histologic examination is illustrated in Figure 1D.5.8.

SUPPORT PROTOCOL 4

Materials Animal cap explants Steinberg’s solution (SS; see recipe) Bouin’s solution: 15 ml picric acid, 5 ml formalin, 1 ml acetic acid, prepare fresh 70% ethanol Xylene Paraffin Delafield’s hematoxylin solution (Sigma, cat. no. 03971) Eosin Y solution (Sigma, cat. no. HT 110216) Canada balsam (Sigma, cat. no. 03984) Special basket, consisting of a glass tube (1 cm × 1 cm) with the bottom covered with a nylon mesh (148-μm grids)

Figure 1D.5.8 Equipment for histological analyses of the differentiation of animal cap explants. The instruments needed for embedding the explants are: (1) special baskets that consist of a glass tube with a nylon mesh on the bottom; (2) watchmaker’s forceps; (3) transfer pipet for handling explants; (4) paraffin molds for embedding the explants in paraffin.

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Paraffin molds 56◦ to 58◦ C paraffin oven Heated wide-bore pipet Microtome Glass microscope slides 45◦ C oven Coverslips Fix animal cap explants 1. Wash animal cap explant samples two times with 2 ml SS. 2. Fix in 1 ml Bouin’s solution for 3 hr. 3. Wash samples with several changes of 70% ethanol to bleach the yellow color of picrate. 4. Dehydrate through a graded series of ethanol (70%, 90%, and 99.5%, in 1-hr incubations). A special basket is used for handling small samples. The samples are placed in the basket, and solution exchanges are performed by simply transferring the basket to the new solution.

Embed in paraffin and section 5. Clear the samples by xylene treatment three times, 15 min each time. 6. Embed the samples in paraffin molds at 56◦ to 58◦ C using a heated wide-bore pipet. 7. Trim the paraffin block for sectioning. 8. Section the samples at 6 μm using a microtome and mount the ribbons of paraffin onto glass slides. Before mounting the paraffin ribbon, place several drops of water onto the slides and then place the ribbons on the water drops.

9. Incubate slides at 45◦ C, to extend the paraffin ribbons, and dry overnight.

Stain with hematoxylin/eosin 10. Deparaffinize the slides with xylene two times, 5 min each time. 11. Hydrate through a graded series of ethanol (99.5%, 90%, 70%, and distilled water, in 5-min incubations). 12. Stain sections with Delafield’s hematoxylin solution for 1 min, wash in running water for over 20 min, and stain with eosin Y solution for 1 min using Coplin jars. 13. Dehydrate the sections with a graded series of ethanol (70%, 90%, and 99.5%, in 5-min incubations). 14. Clear in sections in xylene three times, 5 min each time. 15. Add a coverslip with a drop of Canada balsam. Store slides at room temperature. SUPPORT PROTOCOL 5

Isolation and Differentiation of Xenopus Animal Cap Cells

RT-PCR FOR ANALYZING GENE EXPRESSION IN ANIMAL CAP CELLS To evaluate the tissues and organs induced in animal caps, profiling of marker gene expression is often performed. RT-PCR analysis is very useful and convenient for analyzing quantitatively the expression levels of genes in animal cap explants (Kobayashi et al., 2005).

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Current Protocols in Stem Cell Biology

Materials Animal caps ISOGEN RNA purification reagent (Nippon Gene) Chloroform 2-Propanol 70% (v/v) ethanol, RNase-free RNase-free water Oligo(dT)15 (Roche cat. no. 814-270) 0.1 M DTT dNTP mixture (2.5 mM each) Ribonuclease inhibitor (Takara) Superscript II reverse transcriptase and buffer (Invitrogen cat. no. 18064-022) ExTaq polymerase and 10× ExTaq buffer (Takara cat. no. RR001A) Specific primer sets for detecting target genes (10 pmol/μl each) 200-μl micropipettor 1.5-ml tubes Spectrophotometer 1.5-ml microcentrifuge tubes 42◦ , 60◦ , and 70◦ C heating blocks 200-μl PCR tubes Thermal cycler Extract total RNA from animal cap cells 1. Prepare five to ten animal caps per treatment group for total RNA purification. Generally, 200 to 400 ng of total RNA is obtained per animal cap.

2. Add ISOGEN reagent (100 μl for five caps and 200 μl for ten caps) and homogenize with a 200-μl micropipettor until the cells are completely dissolved. 3. Incubate 5 min at room temperature.

Purify RNA 4. Add chloroform (20 μl for five caps and 40 μl for ten caps) and shake vigorously for 15 sec. 5. Incubate 2 to 3 min at room temperature. 6. Centrifuge 15 min at maximum speed, 4◦ C. 7. Remove the aqueous phase (upper, clear layer) and transfer to a new 1.5-ml tube.

Propanol precipitate RNA 8. Add an equal volume of 2-propanol and mix well. 9. Incubate 10 min at room temperature. 10. Centrifuge 10 min at maximum speed, room temperature. 11. Discard the supernatant (check for precipitate at the bottom of the tube) 12. Add 0.2 to 0.5 ml of 70% ethanol. 13. Centrifuge 3 min at maximum speed, room temperature. 14. Discard all of the liquid (check for precipitate at the bottom of the tube). 15. Dry briefly. 16. Add 1.5 μl of RNase-free water.

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17. Estimate the concentration of RNA using a spectrophotometer.

Synthesize cDNA from total RNA 18. Place the following in a 1.5-ml microcentrifuge tube: 0.5 μg total RNA 1 μl oligo p(dT)15 (400 μg/ml) Distilled water to 8.5 μl. 19. To denature RNA, incubate 5 min in a 60◦ C heating block, and then immediately chill on ice. 20. Add the following solution and mix gently:

4 μl 5× reaction buffer 2 μl 0.1 M DTT 2.5 mM of each dNTP in 4 μl 0.5 μl ribonuclease inhibitor 1 μl Superscript II reverse transciptase. 21. Incubate 1 hr at 42◦ C. 22. Incubate 15 min at 70◦ C to stop the reaction. This mixture can be used for subsequent PCR without additional treatment.

Carry out PCR 23. Add the following items to a 200-μl PCR tube: 1 μl cDNA solution 2 μl 10× ExTaq buffer 1.6 μl dNTP mixture 1 μl forward primer 1 μl reverse primer 13.8 μl distilled water. 24. Add 0.2 μl of ExTaq polymerase, and mix by gently pipetting. 25. Perform PCR. Quantitate PCR results by gel electrophoresis. In general, 25 to 28 cycles of a three-step PCR or 35 to 40 cycles of a two-step (shuttle) PCR are performed; the annealing time, extension time, and the number of cycles are set according to the recommended conditions for each gene. If the PCR products are not efficiently amplified, alternative PCR conditions should be tested, e.g., altering the volume of the cDNA in PCR mixture. In some cases, a decreased (rather than increased) volume of cDNA solution may give better results. Annealing temperature is another important parameter for amplification. If possible, several annealing temperatures should be tested (e.g., using a gradient cycler). Increasing the amount of ExTaq polymerase may also improve the outcome. SUPPORT PROTOCOL 6

Isolation and Differentiation of Xenopus Animal Cap Cells

IMMUNOHISTOCHEMISTRY OF THE INDUCED ANIMAL CAP CELLS Detection of tissue-specific proteins is important for the evaluation of induced animal caps. Immunohistochemistry is the most useful method for protein detection. In contrast to in situ hybridization, immunodetection provides information on the subcellular localizations of marker gene products.

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Materials Induced animal caps Fixation solution: 4% (w/v) paraformaldehyde in PBS (see recipe) 25%, 55%, 75%, and 100% methanol Bleaching solution (see recipe) PBT: 0.1% (v/v) Triton X-100 in PBS (see recipe for PBS) Blocking solution (see recipe) Primary antibody Secondary antibody, alkaline phosphatase (AP)–conjugated AP reaction buffer (see recipe) Color solution: 4.5 μg/ml NBT, 3.5 μg/ml BCIP in AP reaction buffer Screw-cap glass vial Incline shaker Dish Aluminum foil Fluorescent light source Pasteur pipet Prepare fixed caps for immunohistochemistry 1. Transfer induced animal caps to a 5-ml glass vial. 2. Add 1 ml of fixation solution. 3. Place vial on incline shaker for 2 hr at room temperature. 4. Discard fixation solution and add 1 ml methanol. 5. Place vial on incline shaker 5 min at room temperature. 6. Replace methanol with 1 ml of bleaching solution. 7. Place vial on a dish over aluminum foil under a fluorescent light. 8. Incubate ∼5 hr at room temperature until the animal caps are completely bleached. 9. Replace bleaching solution with 1 ml methanol, incubate 5 min, and then replace with >2 ml methanol. In this state, the embryos can be stored for more than 2 months at −20◦ C.

Incubate in primary antibody 10. Transfer animal caps in methanol to a glass vial. 11. Replace solution with 75% (v/v) methanol in water and store 5 min at room temperature. 12. Replace solution with 55% (v/v) methanol in water and store 5 min at room temperature. 13. Replace solution with 25% (v/v) methanol in PBT and store 5 min at room temperature. 14. Replace solution with PBT and incubate 15 min at room temperature. If necessary, add NP-40 to the PBT (final concentration 0.4% v/v) for permeabilization. 15. Replace solution with blocking solution and incubate for at least 30 min at room temperature. 16. Replace solution with PBT containing the appropriate dilution of primary antibody and incubate 2 hr at room temperature or overnight at 4◦ C. Refer to XMMR website

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(http://www.xenbase.org/xenbase/original/www/marker-pages/marker-index.html) for antibodies that work in Xenopus tissue.

Wash and incubate in secondary antibody 17. Replace solution with PBT and place the vial on the incline shaker for 1 hr. 18. Repeat step 17 four times for a total of five times. 19. Replace solution with secondary antibody solution and incubate 2 hr at room temperature or overnight at 4◦ C. In this method, an AP-conjugated secondary antibody is used to recognize the primary antibody. Immunodetection with a HRP-conjugated antibody is also possible.

20. Replace the solution with PBT and place the vial on the incline shaker for 1 hr. 21. Repeat step 20 four times.

Visualize antibody binding 22. Replace solution with AP reaction buffer and incubate 5 min at room temperature. 23. Replace the solution with color solution and incubate in the dark. 24. Check for color development. Typically, color development is done for 1 hr.

25. When the appropriate color appears, replace with fixation solution, which denatures alkaline phosphatase. SUPPORT PROTOCOL 7

WHOLE-MOUNT IN SITU HYBRIDIZATION Whole-mount in situ hybridization (WISH) is a technique that is widely used to study regional mRNA expression. In many studies using animal cap cells, this method facilitates the collection of valuable experimental information. This method is derived from that of Harland (1991). NOTE: All the materials used should be RNase- and DNase-free, and gloves should be worn. The basic regents are prepared according to previously published protocols (Sambrook and Russell, 2001). All materials can be substituted with equivalent items.

Materials

Isolation and Differentiation of Xenopus Animal Cap Cells

Plasmid containing target clone Appropriate restriction enzyme Phenol/chloroform 100% ethanol RNase-free water T3 RNA polymerase (Roche cat. no. 1031163), T7 RNA polymerase (Roche cat. no. 881767), or SP6 RNA polymerase (Roche cat. no. 810274) and 10× transcription buffer Dig RNA labeling mix (Roche cat. no. 1277073) RNase inhibitor (Takara cat. no. 2310A) DNase I (Invitrogen cat. no. 18068-015) Stop solution (see recipe) Hydrolysis buffer (see recipe) 3 M sodium acetate, pH 5.2 MEMFA (see recipe) 50% and 75% ethanol in RNase-free water 25% ethanol in PTw

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PTw (see recipe) 10 μg/ml Proteinase K (see recipe) 0.1 M TEA (see recipe) 4% PFA (see recipe) Hybridization buffer (see recipe) 0.2× and 2× SSC (see recipes) RNase in 2× SSC (see recipe) MAB (see recipe) MAB+BR (see recipe) MAB+BR+SS (see recipe) Anti-digoxigenin-AP, Fab fragment (Roche cat. no. 1093274) AP buffer (see recipe) BM Purple (Roche cat. no. 1442074) 70% and 100% methanol Bleaching solution (see recipe) Spectrophotometer 37◦ and 60◦ C water bath 5-ml screw-cap glass vial Pipet Mild shaker Hybridization incubator 24-well plate Prepare plasmid 1. Linearize plasmid containing target clone by digesting with a suitable restriction enzyme. Check for complete digestion by DNA gel electrophoresis. 2. Phenol/chloroform extract and ethanol precipitate the digested plasmid. 3. Dissolve the digested plasmid in a suitable volume of RNase-free water. Measure the DNA concentration in a spectrophotometer (OD260 ) and adjust to a final concentration of 1 μg/μl.

Label transcripts 4. Set up transcription reaction as follows: 3 μl 1 mg/ml digested plasmid 5 μl 10× transcription buffer 5 μl Dig RNA labeling mix 1 μl RNase inhibitor 2 μl RNA polymerase (SP6, T3, or T7) RNase-free water to 50 μl. 5. Incubate 2 hr at 37◦ C. 6. Check for correct probe synthesis by denaturing gel electrophoresis.

Prepare probe 7. Add 1 μl DNase I and incubate 15 min at 37◦ C. 8. Add 50 μl stop solution and ethanol precipitate. 9. Dissolve in 60 μl hydrolysis buffer on ice. 10. Incubate for the appropriate time period at 60◦ C.

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Synthesized probes are alkali-degraded to final lengths of 300 bp, 200 bp, and 100 bp. Therefore, the appropriate incubation time is calculated using the following formula: Incubation time (min) = (Li – Lf)/0.11 × Li × Lf where Li is the initiation length (kb) and Lf is the final length (kb). A worksheet for this calculation is shown in Table 1D.5.1.

11. Add the final sample (20 μl for three rounds) to 120 μl RNase-free water plus 20 μl of 3 M sodium acetate, pH 5.2, on ice. 12. Ethanol precipitate and dissolve in 40 μl RNase-free water. 13. Check for complete degradation by denaturing gel electrophoresis and measure the concentration (OD260 ). Synthesized probes can be stored up to 6 months at −20◦ C.

Fix animal caps 14. Transfer the treated animal caps to a 5-ml screw-cap vial that is partially (Fig. 1D.5.9A) filled with MEMFA. 15. Gently shake (Video 4) the vial for 1 hr at room temperature. 16. Remove MEMFA and replace with ethanol (Fig. 1D.5.9A). 17. Gently shake (Video 4) the vial for 1 hr at room temperature. 18. Remove ethanol, replace with fresh ethanol, and store at −20◦ C until ready for hybridization. The animal caps can be stored for up to 6 months at −20◦ C.

Perform whole-mount in situ hybridization (see Table 1D.5.2) Day 1 (probe hybridization) 19. Rehydrate the animal caps through an ethanol series (100% ethanol, 75% ethanol in RNase-free water, 50% ethanol in RNase-free water, and 25% ethanol in PTw). Incubate each step for 5 min at room temperature. 20. Wash four times with PTw 5 min each time at room temperature. 21. Treat with 10 μg/ml Proteinase K 1 min at room temperature (2 ml/tube). The timing of this step is crucial. Between steps 21 and 25, the animal caps are fragile, so the solution must be exchanged gently.

22. Wash two times with 0.1 M TEA 1 min each time at room temperature. Table 1D.5.1 Appropriate Incubation Time for Alkali-Degradation of the Synthesized Probes

Time to Lf = 0.3 kbp (min)

Time to Lf = 0.2 kbp (min)

Time to Lf = 0.1 kbp (min)

1.0

21

36

82

0.9

20

35

81

0.8

19

34

80

0.7

17

32

78

0.6

15

30

76

0.5

12

27

73

0.4

8

23

68

Li (kb)

Isolation and Differentiation of Xenopus Animal Cap Cells

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Figure 1D.5.9 Handling of animal cap explants on the whole-mount in situ hybridization. (A) The screw-cap glass vial is partially filled with solution (arrow). (B) Stratified acetic anhydrate diffuses gradually in 0.1 M TEA (see Day 1, Support Protocol 7, step 23).

23. Replace with 4 ml of 0.1 M TEA, stratify with 10 μl acetic anhydride (as in Fig. 1D.5.9B), and allow to stand 5 min at room temperature. If acetic anhydride droplet sinks to the bottom of the vial, then acetylation of animal caps is heterogeneous. Moreover, the animal cap is broken by the direct hit of the droplet.

24. Wash two times with PTw 5 min each time at room temperature. 25. Refix the animal caps in 4% PFA 15 min at room temperature. This step must be timed precisely.

26. Wash five times with PTw 5 min each time at room temperature. 27. Wash with 0.5 ml hybridization buffer 10 min at 60◦ C. 28. Prehybridize in 1 ml hybridization buffer 1 hr at 60◦ C. 29. Hybridize in 1 ml hybridization buffer containing probe (final concentration 1 μg/ml) overnight at 60◦ C.

Day 2 (washing and antibody incubation) 30. Remove the hybridization buffer/probe mix and replace with 1 ml hybridization buffer. Wash 10 min at 60◦ C. 31. Wash three times in 3 ml of 2× SSC 20 min each time at 60◦ C. 32. Replace with 3 ml RNase in 2× SSC and incubate 30 min at 37◦ C. This solution and waste from steps 3 to 5 must be sealed and discarded properly, as they contain high concentrations of RNase.

33. Wash two times in 2× SSC 5 min each time at room temperature. 34. Wash two times in 0.2× SSC 30 min each time at 60◦ C. 35. Wash two times in MAB 10 min each time at room temperature. 36. Wash in MAB+BR 15 min at room temperature. 37. Pre-incubate in 2 ml MAB+BR+SS 1 hr at room temperature. 38. Incubate in 1 ml MAB+BR+SS containing antibody (anti-digoxigenin-AP, Fab fragment) diluted 1:5000 from stock overnight at 4◦ C.

Embryonic and Extraembryonic Stem Cells

1D.5.21 Current Protocols in Stem Cell Biology

Supplement 9

Table 1D.5.2 Worksheet for Whole-Mount In Situ Hybridizationa

Solution

Time

Temperature

Statusb

Day 1 Ethanol

5 min

RT

Swing (roll)

75% ethanol/25% RNase free water

5 min

RT

Swing (roll)

50% ethanol/50% RNase free water

5 min

RT

Swing (roll)

25% ethanol/75% PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

10 μg/ml Proteinase K

1 min

RT

2 ml/stand

0.1 M TEA

1 min

RT

Swing (roll)

0.1 M TEA

1 min

RT

Swing (roll)

0.1 M TEA + acetic anhydrate

5 min

RT

4 ml + 10 μl/stand

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

4% PFA

15 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

PTw

5 min

RT

Swing (roll)

Hybridization buffer

10 min

◦

60 C

0.5 ml/swing (stand)

Hybridization buffer

1 hr

60◦ C

1 ml/swing (stand)

O/N

◦

60 C

1 ml/swing (stand)

10 min

60◦ C

1 ml/swing (stand)

20 min

◦

60 C

3 ml/swing (stand)

20 min

◦

60 C

3 ml/swing (stand)

20 min

◦

60 C

3 ml/swing (stand)

RNase in 2× SSC

30 min

◦

37 C

3 ml/swing (stand)

2× SSC

5 min

RT

3 ml/swing (stand)

2× SSC

5 min

RT

3 ml/swing (stand)

30 min

◦

60 C

3 ml/swing (stand)

0.2× SSC

30 min

◦

60 C

3 ml/swing (stand)

MAB

10 min

RT

Swing (roll)

MAB

10 min

RT

Swing (roll)

Hybridization buffer + probe Day 2 Hybridization buffer 2× SSC 2× SSC 2× SSC

0.2× SSC

Isolation and Differentiation of Xenopus Animal Cap Cells

continued

1D.5.22 Supplement 9

Current Protocols in Stem Cell Biology

Table 1D.5.2 Worksheet for Whole-Mount In Situ Hybridizationa , continued

Solution MAB+BR

Time

Temperature

15 min

RT

Statusb Swing (roll)

MAB+BR+SS

1 hr

RT

2 ml/swing (stand)

MAB+BR+SS+Ab

O/N

4◦ C

1 ml/swing (stand)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

MAB

1 hr

RT

Swing (roll)

AP buffer

5 min

RT

Swing (roll)

AP buffer

5 min

RT

Swing (roll)

Day 3

Coloring solution a Abbreviations: O/N, overnight; RT, room temperature. b Unless indicated otherwise, the vial is partially filled with solution (see Fig. 1D.5.9A). In the case of “swing (roll),”

shake gently as shown in Video 4. The screw-cap glass vials are gently rotated on the low-speed rocking mixer to snake samples thoroughly. In the case of “swing (stand),” shake gently as shown in Video 5. The glass vials are standing and gently rocking on the mixer to shake sample more mildly.

Day 3 (washing and staining) 39. Wash eight times with MAB 1 hr each time at room temperature. 40. Wash two times in AP buffer 5 min each time at room temperature. 41. Transfer the animal caps to a 24-well plate, one vial per well. 42. Replace AP buffer with 1 ml BM Purple, cover with foil, and incubate with rocking until the desired level of staining is achieved. Staining time will vary depending on the level of expression. For example, Xbra mRNA in animal caps treated with 5 ng/ml activin will be detected within 1 hr using this protocol. Although the reaction proceeds more rapidly at room temperature, the embryos tend to show lower background at 4◦ C.

43. Stop the staining reaction by washing thoroughly in MEMFA 2 hr at room temperature. 44. Wash several times with methanol at room temperature. Most of the brown background staining will be removed by these washes.

45. Replace with bleaching solution and bleach until satisfied at room temperature. This step is required for depigmentation of animal cap explants. This step need not be performed in the dark.

46. Wash in 70% methanol 5 min at room temperature. This step is required for depigmentation of animal cap explants.

47. Store in fresh 100% methanol for up to 6 months at 4◦ C.

Embryonic and Extraembryonic Stem Cells

1D.5.23 Current Protocols in Stem Cell Biology

Supplement 9

48. Examine the animal cap for indicating binding of the hybridization probe, shown by the reduction of the pinkish background and clear visibility of a blue signal. After hybridization, embed animal caps in paraffin and section at 10-μm thickness to check internal structures (see Support Protocol 4, without HE-staining).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. To DEPC-treat buffers/water, add 0.05% DEPC, incubate with agitation until completely dissolved, and then autoclave. RNase-free water indicates water that has been treated with DEPC.

AP buffer 100 ml 2× AP buffer (−) (see recipe) 5 ml 2 M MgCl2 1 ml 20% (v/v) Tween 20 94 ml water 0.24 g tetramisole hydrochloride (Sigma cat. no. L9756-5G) Prepare fresh AP buffer (−), 2× 12.11 g Tris 86 ml water Adjust to pH 9.5 with HCl Add 5.84 g NaCl Add water to 500 ml and autoclave Store up to 6 months at room temperature Bleaching solution 2 vol H2 O2 3 vol formamide 15 vol 2× SSC (see recipe) 40 vol water Prepare fresh As H2 O2 is corrosive, toxic, and damaging to skin, appropriate gloves and protective clothing should be worn.

Blocking reagent, 10% 10 g blocking reagent (Roche cat. no. 1096176) 100 ml MAB (see recipe) Store at –20◦ C as a stock solution Heat at 60◦ C and mix to dissolve completely Autoclave Dispense into 10-ml aliquots and store up to 6 months at −20◦ C DeBoer’s solution (DB)

Isolation and Differentiation of Xenopus Animal Cap Cells

110.00 mM NaCl 1.30 mM KCl 0.44 mM CaCl2 3.00 mM HEPES Adjust to pH 7.3 with 1 M NaOH Store up to 6 months at room temperature

1D.5.24 Supplement 9

Current Protocols in Stem Cell Biology

Hybridization solution 50 ml formamide 25 ml 20× SSC (see recipe) 9.5 ml RNase-free water 10 ml 10 mg/ml Torula RNA (Sigma cat. no. R-3629) in RNase-free water 2 ml 10 mg/ml heparin (Sigma cat. no. H3393-100KU) in RNase-free water 0.5 ml 20% (v/v) Tween 20 in RNase-free water 1 ml 10% (w/v) CHAPS (Dojindo cat. no. 349-04722) in RNase-free water 2 ml 0.5 M EDTA Store up to 6 months at −20◦ C Hydrolysis buffer 0.67 g NaHCO3 1.27 g Na2 CO3 Add RNase-free water to 200 ml Check for pH 10 with pH-test paper or a compact pH meter (measure from a single-drop sample) Store up to 3 months at room temperature This solution should not be autoclaved, so prepare with RNase-free reagents and experimental instruments.

MAB 11.61 g maleic acid 8.77 g NaCl 950 ml water Adjust to pH 7.5 with 10 N NaOH Add water to 1 liter Autoclave Store up to 6 months at room temperature MAB+BR 4 vol MAB (see recipe) 1 vol 10% blocking reagent (see recipe) Store up to 6 months at −20◦ C MAB+BR+sheep serum 4 vol MAB+BR (see recipe) 1 vol heat-inactivated sheep serum (Chemicon cat. no. S22-100) Store up to 6 months at −20◦ C Sheep serum is heat-inactivated at 55◦ C for 35 min, dispensed into aliquots, and stored up to 6 months at −20◦ C.

MEM, 10× 104.64 g MOPS 3.80 g EGTA 1.23 g MgSO4 ·12H2 O 300 ml water Adjust to pH 7.4 with 10 N NaOH Add water to 500 ml and autoclave Store up to 6 months at room temperature Embryonic and Extraembryonic Stem Cells

1D.5.25 Current Protocols in Stem Cell Biology

Supplement 9

MEMFA 1 vol 10× MEM (see recipe) 1 vol formaldehyde 8 vol water Prepare fresh This solution is made just prior to use. Since formaldehyde is highly toxic, wear gloves and handle in a chemical hood.

Paraformaldehyde (PFA), 4% 2.4 g paraformaldehyde 12 ml RNase-free water 24 μl 10 N NaOH Heat at 60◦ C, mixing occasionally until completely dissolved Add 48 ml PTw (see recipe) and cool on ice This solution is made just prior to use. As paraformaldehyde is highly toxic, gloves should be worn and handling should be performed in a chemical hood.

PBS, 10× 80 g NaCl 2 g KCl 28.98 g Na2 HPO4 ·12H2 O 2 g KH2 PO4 900 ml water Adjust to pH 7.4 with 10 N NaOH Add water to 1 liter and autoclave Store up to 6 months at room temperature Proteinase K, 10 μg/ml 1 vol 20 mg/ml Proteinase K (Wako cat. no. 163-18131) 2000 vol PTw (see recipe) Store up to 6 months at 4◦ C PTw 100 ml 10× PBS (see recipe) Add water to 1 liter Treat with DEPC Autoclave Add 1 ml Tween 20 and mix well Store up to 3 months at room temperature RNase in 2× SSC 20 vol 10 mg/ml RNaseA (Sigma cat. no. R5000-100MG) in water 1 vol 105 U/ml RNaseT1 (Wako cat. no. 185-01601) in water 10,000 vol 2× SSC (see recipe) Prepare fresh SSC, 0.2×

Isolation and Differentiation of Xenopus Animal Cap Cells

50 ml 2× SSC (see recipe) Add water to 500 ml and autoclave Store up to 6 months at −20◦ C

1D.5.26 Supplement 9

Current Protocols in Stem Cell Biology

SSC, 2× 50 ml 20× SSC (see recipe) Add water to 500 ml and autoclave Store up to 6 months at −20◦ C SSC, 20× 175.3 g NaCl 88.2 g sodium citrate 800 ml water Adjust to pH 7.0 with HCl Add water to 1 liter Treat with DEPC Autoclave Store up to 6 months at room temperature Steinberg’s solution (SS) 58.00 mM NaCl 0.67 mM KCl 0.34 mM Ca(NO3 )2 0.83 mM MgSO4 3.00 mM HEPES 0.01% (w/v) kanamycin sulfate Adjust to pH 7.4 with 1 N NaOH Store up to 6 months at room temperature Stop solution 20 μl 0.1 M NaCl 20 μl 1 M Tris·Cl, pH 7.5 40 μl 0.5 M EDTA, pH 8.0 100 μl 10% (w/v) SDS 820 μl RNase-free water Store up to 3 months at room temperature TEA, 0.1 M 7.5 ml triethanolamine 500 ml RNase-free water 4 ml HCl Store up to 1 month at room temperature COMMENTARY Background Information The animal cap is an excellent tool for analyzing various inductive interactions during early amphibian embryogenesis. It can be induced to differentiate into neural tissue, mesoderm, and endoderm by exposure to specific inducers. For example, in the classical Spemann’s organizer experiment, the blastopore lip was transplanted into the ventral side of a host embryo. The neural tissues of the induced secondary embryo were almost entirely derived from the host ventral ectoderm, which consisted of a part of the animal cap.

In the recombination experiment presented by Nieuwkoop (1969), the animal cap was directly combined with vegetal cells lacking mesoderm cells of the marginal zone; at the end of the culture period, the differentiation of mesodermal tissues was confirmed in the recombinant. This phenomenon is termed mesoderm induction because the mesodermal tissues were induced from the animal cap under the influence of the vegetal endoderm cells. It is this pluripotency that makes animal cap cells the amphibian equivalent of embryonic stem cells.

Embryonic and Extraembryonic Stem Cells

1D.5.27 Current Protocols in Stem Cell Biology

Supplement 9

Isolation and Differentiation of Xenopus Animal Cap Cells

In the late 1980s, the pluripotency of animal caps enabled remarkable advances in studies of mesoderm-inducing factors. Several peptide growth factors belonging to the fibroblast growth factor (FGF) and transforming growth factor-β (TGF-β) families were revealed to be capable of inducing mesodermal tissue formation from animal caps (reviewed in Asashima et al., 2008). One of the later molecules, activin, induces almost all mesodermal tissues in a dose-dependent manner (Green and Smith, 1990; Ariizumi et al., 1991a,b; Green et al., 1992). Moreover, activin in combination with other molecules can induce the formation of multiple organs in animal caps. For example, pronephros (Moriya et al., 1993) and pancreas (Moriya et al., 2000) are induced in animal caps treated with a combination of activin and retinoic acid (see Alternate Protocol 1). The most characteristic property of activin is the induction of organizer activity in animal caps. Following treatment with a high concentration of activin (100 ng/ml), the animal cap induces a secondary embryo, as does the Spemann’s organizer when transplanted into another embryo. It is possible to control organogenesis and to design a fundamental embryonic body plan using activin as the inducer and the animal cap as the reacting tissue (Ariizumi and Asashima, 1994). The range of utility of the animal cap is extended by combining it with the microinjection method (see Basic Protocol 2). Animal caps obtained from mRNA- or DNAinjected embryos provide much information about the function of the target gene. Investigators can analyze changes in competency or reactivity by comparing these animal caps treated with a specific inducing molecule with non-injected animal caps. The microinjection technique is also applied in cell-lineage tracing experiments, such as the in vivo transplantation of in vitro–induced animal caps. For example, the tissues or organs derived from the transplanted animal caps can be detected in the host embryos if the caps are derived from embryos injected with a fluorescent dye or a gene that encodes an enzyme (e.g., β-gal, HRP) at the early cleavage stages. Depending on the purpose of the experiment, the researcher may be expected to combine the animal cap assay with the microinjection technique. The animal cap assay in conjunction with several methods for analyzing the differentiation of animal caps at the histologic and molecular levels have been described in this unit. Excellent guide books, such as Kay and Peng (1991) and Sive et al.

(2000), provide more detailed descriptions of these protocols.

Critical Parameters In the animal cap assay, the isolated caps form irregular-shaped epidermis (atypical epidermis) in the absence of inducers but can be induced to form neural, mesodermal, and endodermal tissues by the addition of certain inducers in a saline solution. The differentiation of notochord and muscle in the animal cap explants indicates a mesoderm-inducing activity. If the saline solution contains a neural inducer, archencephalic structures, such as the forebrain and eyes, will be induced in the explants. The utmost care must be taken when identifying the neural inducer, since animal caps are susceptible to artificial stimulation. For example, animal caps cultured in a highsalt solution (>100 mM NaCl) sometimes form neural tissues in the absence of inducers. To obtain reliable results for the animal cap assay, experimenters should pay close attention to the following parameters. First, although animal caps are competent up to stage 10 (early gastrula), their responses to inducers are slightly different. The choice of cap age is dependent upon the desired outcomes; thus, accurate staging of embryos is important. The late blastula (stage 9) is used as the standard for the animal cap assay in the authors’ laboratory. Second, concerns arise regarding the size of the animal caps dissected. Any size is acceptable as long as the control animal cap forms atypical epidermis in the absence of inducer. A large animal cap may be contaminated with marginal zone cells, which can differentiate autonomously into mesodermal tissues. In the authors’ experience, the most reliable animal cap size is 0.5 mm × 0.5 mm. Third, the duration of exposure of animal caps to the inducer also influences their differentiation patterns. For example, a brief exposure (5 min) to 10 ng/ml activin causes the differentiation of ventral mesoderm, such as mesenchyme and mesothelium, while a long exposure (3 hr) to the same dosage leads to muscle differentiation in animal caps (Ariizumi et al., 1991a). The developmental stage of the animal cap is also very important to the success of heart induction in vitro. When the animal caps are obtained from embryos at stage 9 or later, it is difficult to induce a beating heart in the dissociation/reaggregation system (see Alternate Protocol 2). The number of cells in the reaggregate also affects the efficacy of heart formation. The frequency of heart formation is 80% to 100% when five animal caps (∼1000 cells)

1D.5.28 Supplement 9

Current Protocols in Stem Cell Biology

Table 1D.5.3 Troubleshooting Guide for Animal Cap Assay

Problem

Possible cause

Solution

Animal cap does not survive Weak or abnormal eggs and embryos Obtain highest quality fertilized eggs Excessive dejellying

Wash the dejellied embryos as quickly as possible with a large volume of saline

Bacterial contamination

Autoclave the saline and add antibiotics, such as kanamycin sulfate (0.1 mg/ml), to the saline

Improper temperature

All operations and culturing of the animal cap explants should be performed at 20◦ to 22◦ C

Density of animal caps in the culture Fewer than ten caps per 1 ml of test solution or dish or plate is too high culture medium is reasonable The concentration of inducing factor Adjust concentration of inducer is too high Animal cap curls up too Improper temperature rapidly after dissection from the embryo

Animal cap cells disperse and adhere to the dish

Weak effect of the inducer on the animal caps

Improper concentration of NaCl in the saline

It is possible to delay animal cap curling by increasing the concentration of NaCl from 60 mM to 90 mM

Incorrect composition of the saline solution

Check the calcium ion concentration of the saline solution

BSA is not included in the saline solution

Add 0.1% BSA to the saline solution, to prevent cells adhering to the dish

Deactivation of the inducer

Avoid freezing and thawing the inducer

BSA is not added to the saline solution

Add 0.1% BSA to the test solution to avoid the adsorption of inducer to the dish

Inappropriate concentration and duration of treatment

Adjust the concentration and duration of treatment

Low competency of the animal cap

Select embryos of the appropriate stage

Differentiation of mesoderm Contamination of the animal cap and/or endoderm in the with marginal zone cells absence of inducer

Differentiation of neural tissue in the absence of inducer

Lower the temperature to 16◦ to 18◦ C

Trim the caps to remove marginal zone cells

Contamination of the animal cap with yolky endoderm cells

Remove any vegetal yolky cells, which are large and white compared with the animal cap cells

Inappropriate formulation of the saline solution

Lower the concentration of NaCl in the saline solution to 60 mM. Animal cap cells differentiate neural tissue autonomously upon exposure to >100 mM NaCl.

Contamination of the animal cap with marginal zone cells

Trim the caps and remove marginal zone cells, which may induce neural tissues as a secondary induction event

Embryonic and Extraembryonic Stem Cells

1D.5.29 Current Protocols in Stem Cell Biology

Supplement 9

are contained in a single reaggregate (Ariizumi et al., 2003).

Troubleshooting See Table 1D.5.3 for troubleshooting suggestions for the animal cap assay.

RT-PCR within a few hours of the initiation of induction. Typical differentiation patterns can be observed in the histologic sections of animal cap explants that are cultured for >2 days at 20◦ C.

Literature Cited Anticipated Results When activin is used as an inducer in the animal cap assay, its effect on the caps is distinctly dose-dependent, with induction of more dorsal mesoderm as the concentration increases. The activin-treated animal caps show rounding up within 3 hr of the initiation of treatment. They form spheres with the original blastocoel surface in the interior and they begin to elongate after ∼3 hr. The degree of elongation depends on the concentration of activin used. Excessive elongation is observed for caps treated with 5 to 10 ng/ml of activin (see Video 6). This phenomenon is considered to mimic the convergent extension of dorsal mesoderm during gastrulation in normal development. At the end of the culture period (2 to 3 days), the animal cap explants show obvious histodifferentiation patterns (Ariizumi et al., 1991b). Activin concentrations of 0.5 to 1 ng/ml result in the differentiation of ventral mesoderm, such as blood cells, mesothelium, and mesenchyme. Muscle is formed at 5 to 10 ng/ml activin, and the notochord, which is the most dorsal mesoderm, is induced at 50 to 100 ng/ml activin. The expression of tissuespecific genes is detected in activin-treated animal caps in the same manner as in normal development. The dissociation/reaggregation protocol (see Alternate Protocol 2) synchronizes the response of animal cap cells. Dissociated cells can be exposed to a more uniform concentration of inducer when compared to the multilayered animal caps. By using the dissociation/reaggregation protocol, the dosedependent mesoderm induction of activin can be observed with clearer dose thresholds (Green and Smith, 1990; Green et al., 1992).

Time Considerations

Isolation and Differentiation of Xenopus Animal Cap Cells

Removal of animal caps should be completed within 1 to 2 hr. The experimenter must excise the animal caps from the embryos as quickly as possible, to avoid variability in the developmental stages of the caps. It is possible to continue the manipulations over several hours when embryos are generated through successive rounds of in vitro fertilization at appropriate intervals. The gene expression patterns of the animal caps can be detected by

Ariizumi, T. and Asashima, M. 1994. In vitro control of the embryonic form of Xenopus laevis by activin A: Time and dose-dependent inducing properties of activin-treated ectoderm. Develop. Growth Differ. 36:499-507. Ariizumi, T., Sawamura, K., Uchiyama, H., and Asashima, M. 1991a. Dose- and time-dependent mesoderm induction and outgrowth formation by activin A in Xenopus laevis. Int. J. Dev. Biol. 35:407-414. Ariizumi, T., Moriya, N., Uchiyama, H., and Asashima, M. 1991b. Concentration-dependent inducing activity of activin A. Roux’s Arch. Dev. Biol. 200:230-233. Ariizumi, T., Kinoshita, M., Yokota, C., Takano, K., Fukuda, K., Moriyama, N., Malacinski, G.M., and Asashima, M. 2003. Amphibian in vitro heart induction: A simple and reliable model for the study of vertebrate cardiac development. Int. J. Dev. Biol. 47:405-410. Asashima, M., Michiue, T., and Kurisaki, A. 2008. Elucidation of the role of activin in organogenesis using a multiple organ induction system with amphibian and mouse undifferentiated cells in vitro. Develop. Growth Differ. 50:S35-S45. Green, J.B. and Smith, J.C. 1990. Graded changes in dose of a Xenopus activin A homologue elicit stepwise transitions in embryonic cell fate. Nature 347:337-338. Green, J.B., New, H.V., and Smith, J.C. 1992. Responses of embryonic Xenopus cells to activin and FGF are separated by multiple dose thresholds and correspond to distinct axes of the mesoderm. Cell 71:731-739. Harland, R.M. 1991. In situ hybridization: An improved whole-mount method for Xenopus embryos. Methods Cell Biol. 36:685-695. Kay, B.K. and Peng, H.B., eds. 1991. Methods in Cell Biology. Xenopus laevis: Practical Use in Cell and Molecular Biology. Academic Press, San Diego, California. Kobayashi, H., Michiue, T., Yukita, A., Danno, H., Sakurai, K., Fukui, A., Kikuchi, A., and Asashima, M. 2005. Novel Daple-like protein positively regulates both the Wnt/beta-catenin pathway and the Wnt/JNK pathway in Xenopus. Mech. Dev. 122:1138-1153. Moriya, H., Uchiyama, H., and Asashima, M. 1993. Induction of pronephric tubules by activin and retinoic acid in presumptive ectoderm of Xenopus laevis. Develop. Growth Differ. 35:123128. Moriya, N., Komazaki, S., Takahashi, S., Yokota, C., and Asashima, M. 2000. In vitro pancreas formation from Xenopus ectoderm treated with activin and retinoic acid. Develop. Growth Differ. 42:593-602.

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Current Protocols in Stem Cell Biology

Nieuwkoop, P.D. 1969. The formation of mesoderm in urodelan amphibians, Pt. 1: Induction by the endoderm. Roux’ Arch. Entwicklungsmech. Org. 162:341-373. Nieuwkoop, P.D. and Faber, J. 1967. Normal Table of Xenopus laevis (Daudin). NorthHolland Publishing, Amsterdam. Sambrook, J. and Russell, D.W. 2001. Molecular Cloning. Cold Spring Harbor Laboratory Press. Cold Spring Harbor, New York. Sive, H.L., Grainger, R.M., and Harland, R.M. 2000. Early development of Xenopus laevis. Cold Spring Harbor Laboratory Press. Cold Spring Harbor, New York.

Embryonic and Extraembryonic Stem Cells

1D.5.31 Current Protocols in Stem Cell Biology

Supplement 9

SECTION 1E Isolation of Stem Cells from Extraembryonic Tissues INTRODUCTION his section focuses on methods for obtaining stem cells from the extraembryonic membranes and, more specifically, the placenta and umbilical cord. Compared to human and nonhuman primate embryos, little is known about the nature of progenitor cells that are harbored within the placenta and its associated extraembryonic structures (e.g., the amnion, the fluid it produces, and the umbilical cord). However, there is a great deal of interest in interrogating this compartment because the component cells, either embryonic or fetal depending on the gestational age of the tissue, could be an important source of stem progenitors. The differentiative capacity of these cells also awaits investigation. For example, we do not know whether primate extraembryonic stem cells have the apparently irreversible lineage restrictions that are imposed during the early stage of mouse development or whether they retain more plasticity, which in turn would greatly expand their utility as both research and clinical tools.

T

The contributions to this section provide insights into these outstanding questions. At one end of the spectrum, UNIT 1E.1 describes a method for isolating a subpopulation of placental cells that can be directed toward a hepatocyte fate. This surprising finding suggests possible differences in the molecules basis of embryonic and extraembryonic lineage restriction in mice and humans. UNIT 1E.2 describes methods for producing stem cells from amniotic fluid and placenta. In summary, it is very likely that the extraembryonic tissues are an interesting source of many different progenitor populations. Of note is the fact that they are routinely discarded after birth. Thus, compared to cells obtained from the embryo or fetus proper, fewer regulatory issues are involved in studies of cells isolated from the amnion/chorion, making the extraembryonic tissues a source of human progenitors that is routinely and widely available to the research community. Nevertheless, we note that the same institutional approvals and HIPPA regulations that are required for work with other tissues apply here as well. Susan J. Fisher

Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1E.0.1 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e00s1 C 2007 John Wiley & Sons, Inc. Copyright


1E.0.1 Supplement 1

Isolation of Human Placenta-Derived Multipotent Cells and In Vitro Differentiation into Hepatocyte-Like Cells

UNIT 1E.1

Hsing-I Huang1 1

Cathay General Hospital, Taipei, Taiwan

ABSTRACT Several types of progenitor cells can be isolated from various human adult tissues such as bone marrow, adipose tissues, and umbilical cord. Placental tissue collected after labor and delivery can provide a valuable source for adult stem cells. These progenitor cells, termed placenta-derived multipotent cells (PDMCs), are fibroblast-like cells which can attach on the bottom of culture vessels. PDMCs are capable of differentiating into various cells such as adipocytes, osteoblasts, chondrocytes, and neurons. Recently, we showed that PDMCs also possess the ability to differentiate into hepatocyte-like cells. This unit describes the protocols for isolation of PDMCs from human term placental tissue and for setting up in vitro differentiation of PDMCs toward hepatocyte-like cells. These cells not only express the characteristics of human liver cells, but also demonstrate several C 2007 functions of typical hepatocytes. Curr. Protoc. Stem Cell Biol. 1:1E.1.1-1E.1.9.
by John Wiley & Sons, Inc. Keywords: placenta r hepatocytes r differentiation r isolation r multipotent progenitors

INTRODUCTION This unit presents procedures for isolation of placenta-derived multipotent cells (PDMCs; Fig. 1E.1.1) from human placental tissues and a protocol for in vitro differentiation of these cells into hepatic cells. The first protocol (see Basic Protocol 1) presents a method for isolation of the progenitor cells from term placenta. Human term placenta should be kept sterile and processed no later than 24 hr after the delivery. The placental tissue is then minced to small pieces. After treatment with trypsin/EDTA, the freed cells are washed and then seeded on culture vessels. The critical parts of successful isolation include keeping the tissue and cells clear of bacterial or fungal contamination and keeping the tissue cells alive. Once the tissues are dried or fixed in fixative solution, they are not appropriate materials for culture. This unit also describes a method that allows the induction of differentiation of isolated PDMCs toward hepatocyte-like cells (see Basic Protocol 2). Expanded PDMCs are seeded on poly-L-lysine-coated plates and treated with defined medium. A change in cellular morphology from fibroblast-like to polygonal epithelial-like can be observed within 7 days of treatment. Critical to the success of this protocol are the coating of culture surfaces and the growth factors used to stimulate the differentiation. However, after the differentiation, these cells lose their proliferation capacity; thus, the cell numbers will not increase with continued cultivation. The protocols in this unit work for human placental tissues but not for mouse placenta. In addition, the procedures should not be used for processing other human fetal tissues such as amniotic membrane. PDMCs cannot be isolated from every placental tissue sample. However, keeping the tissue sterile and carefully handling it can increase the rate of successful PDMC isolation to ∼50%. Extraembryonic Lineages Current Protocols in Stem Cell Biology 1E.1.1-1E.1.9 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e01s1 C 2007 John Wiley & Sons, Inc. Copyright


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Figure 1E.1.1 Fibroblast-like cells appear on culture vessels 10 days after the first seeding of placental cells. Medium was changed on day 7.

NOTE: Ethics approval required. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

ISOLATION OF PDMCs FROM HUMAN PLACENTA This protocol describes a simple method for isolation of PDMCs from human placental tissue. Human placenta contains various cell populations including trophoblasts, epithelial cells, and some blood cells. However, most of these cells are incapable of attachment and proliferation under these culture conditions. After cultivation for 2 weeks, epithelial cells and fibroblast-like cells will appear as colonies. Finally, only the fibroblast-like cells can keep dividing. The epithelial cells will constitute 10% to 20% of the culture; the fibroblasts will not overgrow.

Materials

Isolation of PDMC from Human Placenta

Donor for term placenta Expansion medium (see recipe), prewarmed before use Dulbecco’s phosphate-buffered saline without calcium or magnesium (CMF-DPBS; Invitrogen, cat. no. 21600) 70% (v/v) ethanol Trypsin/EDTA solution: 0.5% (w/v) trypsin/0.5 mM EDTA (Invitrogen, cat. no. 15400)

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100-mm culture dishes 15-ml polypropylene centrifuge tubes Tweezers, sterile Scissors, sterile Centrifuge 25-cm2 tissue culture flasks Inverted microscope Additional reagents and equipment for counting cells (Phelan, 2006) Collect and wash placental tissue 1. Collect placental tissue samples immediately after delivery. Place each sample in 10 to 20 vol sterile cold expansion medium (∼1:1 ratio of tissue to medium) and place in a transport container. Keep cold and transport to the laboratory as soon as possible (<24 hr). The author uses the part of the placenta near the umbilical cord. Placental tissue is collected from the fetal side of the placenta. PDMC will be verified by immunophenotyping of the appropriate markers (see Commentary). Term placenta is full of blood (maternal and fetal) that has to be removed by extensive washing in a large volume of CMF-DPBS or culture medium. At least 1000 ml of washing solution is needed per 100 g of tissue (wet weight) to remove trapped blood cells (see step 2).

2. Rinse placental tissue sample eight to ten times, each time with 25 ml CMF-DPBS, to remove trapped blood cells. Rinse the tissue sample briefly with 5 ml 70% ethanol, then twice, each time with 10 ml CMF-DPBS, and put in a 100-mm culture dish. This step can be performed repeatedly if necessary.

3. For a 7-g sample, add 7 ml of expansion medium to the culture dish to keep the sample moist. Dried-out tissue samples cannot be used for the cell culture experiments because many cells are dead.

4. Remove the amniotic membrane. Use tweezers and scissors to mince the placental tissues into small pieces (<0.2 cm3 ). In the author’s laboratory, PCR analysis to detect SRY (see Commentary) is performed to confirm that placental tissue is of fetal origin.

5. Decant the culture medium. 6. Apply 10 ml CMF-DPBS to wash the minced tissues, and remove the medium. This procedure is to remove the medium to improve the effects of trypsin/EDTA digestion. Thus, this step can be repeated several times.

Digest sample 7. For 3 g tissue, add 3 ml of trypsin/EDTA solution to the culture dish so as to completely cover the cells. Incubate 10 min. The incubation time should not exceed 30 min because it could harm or the cells or cause cell death

8. Add 7 ml expansion medium to the 100-mm culture dish to stop the trypsin/EDTA reaction. 9. Transfer the entire suspension from the culture dish to a 15-ml centrifuge tube.

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10. Centrifuge 5 min at 258 × g, room temperature. Decant the supernatant. Resuspend pellet in 10 ml of expansion medium.

Seed cultures 11. Seed the cells on 100-mm tissue culture dishes or 25-cm2 tissue culture flasks (poly-L-lysine treatment not needed). Begin incubation. 12. At day 2 and day 3, check by microscopy for signs of bacterial or fungal contamination. If contamination occurs, do not open the caps; dispose of the infected culture vessels immediately.

Expand cultures 13. After incubation for 7 days, check for the appearance of attached cells on the culture surfaces using an inverted microscope. Remove unattached cells and old medium by aspirating the medium, and replace with 10 ml fresh expansion medium. From this point on, change the medium every 7 days. Sometimes colonies can be seen at this time point.

14. After another 7 days, check for the appearance of attached cells on the culture surfaces using an inverted microscope. At this time, several cell colonies could be observed. More attached cells should be seen than at step 13.

15. Subculture the cells as follows when they reach 90% confluency: a. b. c. d. e. f. BASIC PROTOCOL 2

Remove culture medium and wash attached cells with CMF-DPBS. Add 3 ml trypsin/EDTA solution and incubate 5 min at 37◦ C. Add 10 ml expansion medium to inactivate trypsin. Transfer cell suspension to a 15-ml conical centrifuge tube. Centrifuge 5 min at 258 × g, room temperature. Remove supernatant. Resuspend cells in fresh culture medium and replate in fresh culture dishes.

HEPATIC DIFFERENTIATION OF PDMCS IN VITRO This protocol describes the steps for setting up a differentiation experiment. PDMCs can be induced to differentiate into hepatocyte-like cells, which express liver cell–specific markers and show some bioactivities restricted to hepatocytes.

Materials 0.01% (w/v) poly-L-lysine solution (mol. wt. 70,000 to 150,000; Sigma-Aldrich), filter sterilized Dulbecco’s phosphate-buffered saline without calcium or magnesium (CMF-DPBS; Invitrogen, cat. no. 21600) Culture of PDMCs, 80% to 90% confluent (Basic Protocol 1) Trypsin/EDTA solution: 0.5% trypsin/0.5% mM EDTA (Invitrogen, cat. no. 15400) Expansion medium (see recipe) Medium A (see recipe) Medium B (see recipe)

Isolation of PDMC from Human Placenta

6-well tissue culture plates or 35-mm tissue culture dishes 15-ml centrifuge tubes Centrifuge Additional reagents and equipment for counting viable cell (Phelan, 2006)

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Prepare culture vessel 1. Cover the inner surfaces of culture vessels with 0.01% poly-L-lysine solution and incubate 30 min at room temperature. There are several reports of experiments using fibronectin as coating material for induction of hepatic differentiation; however, fibronectin does not work with PDMCs.

2. Withdraw coating solution from culture vessels using pipets. 3. Wash the coated surfaces twice, each with 2 ml CMF-DPBS twice. 4. Air dry the culture dish in a sterile tissue culture hood.

Collect PDMCs 5. Harvest PDMCs from culture dish by treating the attached cells with 3 ml of trypsin/EDTA solution for 5 min at 37◦ C. Decant the trypsin/EDTA solution. 6. Add 7 ml 37◦ C expansion medium to the culture dish to stop the activity of trypsin. Collect PDMCs by pipetting culture medium over the surface of the plate. 7. Transfer the PDMCs and medium to a 15-ml centrifuge tube. 8. Centrifuge 5 min at 179 × g, room temperature. Aspirate the supernatant. 9. Wash the harvested PDMCs three times, each time by adding 10 ml expansion medium, centrifuging 5 min at 179 × g, room temperature, and removing the supernatant and resuspend the pellet in 10 ml expansion medium. 10. Count the number of viable harvested PDMCs using trypan blue exclusion. 11. Adjust the cell concentration to 1× 106 cells/ml by adding expansion medium.

Establish differentiation cultures 12. Seed the PDMCs at a concentration of 1.7 × 104 cells/cm2 on poly-L-lysine-coated culture vessels. The seeding concentration is critical for setting up the differentiation experiments.

13. Incubate 18 to 24 hr. 14. After incubation, carefully remove the culture medium from the culture vessels, wash the attached cells with 2 ml CMF-DPBS, then add 2 ml medium A to replace the decanted culture medium. The wash procedure should be performed gently to avoid the loss of attached cells. Because PDMCs do not adhere tightly to the dish, rapid movements can cause the detachment of these cells.

15. Incubate another 12 to16 hr.

Induce and monitor differentiation 16. Remove the medium A and wash the PDMCs twice, each time with 2 ml CMF-DPBS. 17. Add 2 ml medium B the dish to induce cell differentiation. 18. Incubate cultures. Observe and check the differentiation of cells frequently. The changes of cell morphology from fibroblast-like to epithelial-like can be observed 3 to 5 days after the addition of medium B. The author has applied immunohistochemical staining with anti-HepPar-1 (Dako) to identify differentiated cells. Extraembryonic Lineages

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. for suppliers, see SUPPLIERS APPENDIX.

Expansion medium Dulbecco’s Modified Eagle Medium, high-glucose formulation (DMEM-HG) supplemented with: 10% (v/v) fetal bovine serum (FBS) 100 U/ml penicillin 100 µg/ml streptomycin 3.7 g/liter NaHCO3 Store up to 1 month at 4◦ C Medium A Mixture of 60% (v/v) Dulbecco’s Modified Eagle Medium, low-glucose formulation (DMEM-LG) and 40% (v/v) MCDB 201 medium (Sigma-Aldrich), supplemented with the following: 1× insulin-transferrin-selenium (ITS; Sigma, cat. no. I3146 or Invitrogen) supplement (add from 100× stock) 4.7 µg/ml linoleic acid 1× (1 mg/ml) bovine serum albumin (BSA, Cohn Fraction V) 10–9 M dexamethasone 10–4 M L-ascorbic acid 2-phosphate (Sigma) 100 U/ml penicillin 100 µg/ml streptomycin 2% (v/v) fetal bovine serum (FBS) 10 ng/ml recombinant human epidermal growth factor (Invitrogen) 10 ng/ml recombinant human PDGF-BB (R&D Systems) Store up to 2 weeks at 4◦ C Medium B Mixture of 60% (v/v) Dulbecco’s Modified Eagle Medium, low-glucose formulation (DMEM-LG) and 40% (v/v) MCDB 201 medium (Sigma-Aldrich), supplemented with the following: 1× insulin-transferrin-selenium (ITS; Sigma, cat. no. I3146 or Invitrogen) supplement (add from 100× stock) 4.7 µg/ml linoleic acid 1× (1 mg/ml) bovine serum albumin (BSA, Cohn Fraction V) 10–9 M dexamethasone 10–4 M L-ascorbic acid 2-phosphate (Sigma) 100 U/ml penicillin 100 µg/ml streptomycin 10 ng/ml human fibroblast growth factor 4 (Sigma) 20 ng/ml recombinant human hepatocyte growth factor (Peprotech) Store up to 2 weeks at 4◦ C COMMENTARY Background Information

Isolation of PDMC from Human Placenta

Stem cells are known to be capable of differentiating toward various types of somatic cells. However, stem cells isolated from the different origins have varying differentiation potential. Usually, the stem cells isolated from

embryonic tissue such as the blastocyst have been considered to have superior differentiation abilities in comparison with stem cells isolated from adult tissues. However, recent research provides evidence that several adult stem cells, the progenitors isolated from adult

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tissues, are able to “transdifferentiate” toward cells that belong to other lineages. The transdifferentiation potential of adult stem cells thus opens the door to applying these cells in future medical therapy. Recently, the author’s laboratory and other groups reported that placental tissue could serve as a source for isolation of progenitor cells (In’t Anker et al., 2004; Yen et al., 2005). These cells, termed placenta-derived multipotent cells (PDMCs), have fibroblastlike morphologies and multilineage differentiation ability. PDMCs share some properties with bone marrow-derived mesenchymal stem cells (BM-MSCs), including the expression of CD105, CD29, CD44, CD90, SH3, and SH4, and the ability to differentiate into mesodermal lineage cells. However, PDMCs are negative for STRO-1, which is a marker found on BMMSCs. In addition, PDMCs possess differentiating abilities comparable to BM-MSCs. PDMCs have been demonstrated to be able to differentiate into cells from all three germ layers (Yen et al., 2005; Chien et al., 2006). In comparison with bone marrow, placental tissue has the advantages of being an easily accessible and relatively young tissue. It appears that placenta may serve as an attractive source of progenitor cells. One of the advantages of these protocols is easy handling; the tissue processed in the author’s laboratory is usually no more than 8 cm × 2 cm × 1 cm from the area near the umbilical cord, which is smaller than the amount used for some other protocols. The time needed for processing is short when compared to protocols that take several hours for enzymatic digestion. Once PDMCs are isolated, they can be easily propagated and characterized according to their expression of specific immunophenotypes or by means of differentiation assays for the various types of cells. PDMCs can be frozen and preserved in liquid nitrogen. According to the author’s experience, thawed PDMCs express the same properties as the cells before they have been frozen including the ability to differentiate into the same types of cells to the same extent. The liver has long been considered an organ that is capable of repairing itself. Liver stem cells, which are located within the canals of Hering, can differentiate into parenchymal hepatocytes and bile ductular cells (Evarts et al., 1987). Recent findings demonstrate that adult stem cells isolated from non-liver tissues such as bone marrow, umbilical cord blood, and human islet can be induced to differenti-

ate into hepatocyte-like cells (Petersen et al., 1999; Kakinuma et al., 2003; von Mach et al., 2004). Furthermore, some of the differentiated cells not only express hepatocyte-specific markers (CD105, CD29, CD44, CD90, SH3, and SH4) but also possess liver cell bioactivities, including urea production, albumin secretion, and glycogen storage (Schwartz et al., 2002; Lee et al., 2004). In addition to the similarities to BMMSCs, PDMCs are positive for alpha fetoprotein (AFP) and c-Met expression (Chien et al., 2006). AFP and c-Met are markers of hepatic stem cells in the early-stage human embryo (Kubota et al., 2002; Suzuki et al., 2004). Based on these characteristics, the author’s group tried setting up hepatic differentiation experiments for PDMCs. Using the method described in Basic Protocol 2, results were obtained suggesting that differentiated PDMCs show hepatocyte-like morphology and express liver cell–specific markers as assessed by immunocytochemical staining using anti-hepatocyte antibody (OCH1E5; Dako; Kakinuma et al., 2003). Bioactivity assays also indicate that PDMC-derived hepatocyte-like cells can take up lipoprotein and even store glycogen (Chien et al., 2006). These results are similar to those obtained using BM-MSCs as the progenitors.

Critical Parameters and Troubleshooting It is essential to test the batch of fetal bovine serum, based on growth assays of PDMC and maintenance of the undifferentiated state, before setting up the experiments. Some batches of serum work well in PDMC culture while other batches could induce the senescence of progenitor cell cultures. However, increasing the percentage of serum in expansion medium seems not to yield any difference in cell growth rate. Medium containing 10% serum is sufficient for propagation. The freshness of collected placental tissues is another factor that can affect successful isolation of PDMCs. The placenta samples should be processed within 24 hr after delivery. During transport and storage, keep the tissue in 4◦ C refrigerator, especially if the sample has to wait more than 24 hr to be processed. The tissue should be handled aseptically to minimize the possibility of bacterial or fungal contamination. That is the reason why the author prefers to rinse the tissue with alcohol before processing it; cell toxicity is minimized by keeping ethanol exposure short and

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Isolation of PDMC from Human Placenta

immediately washing with DPBS following the ethanol rinse. Placental tissues contain various blood cells, including red blood cells and some leukocytes. Some blood cells remain in the cultures; some of these are unattached cells and can easily be removed when the medium is exchanged, while others are attached to the plate bottom. These latter cells attach tightly to the culture surface and are not released by trypsin treatment; therefore, they can be separated from PDMCs. In some protocols describing the isolation of MSCs, the seeding concentration seems to be an important factor for successful isolation of stem cells. According to the authors’ experience, the initial seeding density is not a critical factor in isolation of PDMCs. However, plating at excessively low densities (<105 cells/cm2 ) may yield no colony formation. For the hepatic differentiation assay, the coating of the culture surfaces makes a difference. The type of coating material plays an important factor in hepatic differentiation of PDMCs. In the author’s laboratory, polyL-lysine has proven effective in induction of the differentiation, while fibronectin has been shown to be ineffective. However, the mechanism by which the coating reagent exerts its effects on cell differentiation is not yet understood. The other factors that affect differentiation are the growth factors and reagents added to medium B (see Reagents and Solutions), which is used to induce differentiation. The effects that these molecules have on cells might be through cell surface receptors, or they may produce changes in the intracellular concentrations of some specific ions. They may activate or inactivate a specific transduction pathway, thus driving the target cells to convert to specific types of cells. Hepatic growth factor (HGF) and fibroblast growth factor 4 (FGF-4) are effective growth factors in hepatic differentiation of PDMCs as well as bone marrow–derived MSCs. However, some reagents, such as DMSO, an effective inducer for progenitor cells such as adipose stromal cells (Seo et al., 2005), are toxic to PDMCs. When no cells attach to culture dishes or flasks, there are several possible explanations. First, the placental tissue might not be fresh enough, so the native progenitor cells die when the tissue sample is waiting for processing. Second, the seeding concentration might be too low. Third, the composition of medium might be altered. One of the most critical factors in isolation of PDMCs is the medium. The

serum must be checked before use to ensure that it supports the growth of PDMCs. Several factors can affect the outcome of the differentiation experiment. These factors include the passage of cells used (preferably passage numbers 4 to 15) and the varied differentiation abilities of each clone. Try using more placentas and using cells in their early passages to help obtain good differentiation results.

Anticipated Results When successfully isolated, colonies containing 10 to 50 fibroblast-like PDMCs will appear on the surface of culture dishes after 2 to 3 weeks of culture. The morphology of these cells is long and thin. They can be easily identified by their appearance when compared to other attached cells, such as the multinucleated trophoblasts and the polygonal epithelioid cells. However, because they are easily propagated and removed by trypsin treatment, a homogeneous population of PDMCs can be attained after subculturing these cells several times (three to four passages). For the hepatic differentiation experiments, a change in cell morphology can be observed after 3 to 5 days treatment with medium B. The expression of some hepatocyte-specific markers can be detected as early as day 7.

Time Considerations The time required for isolation of PDMCs depends on how many placental tissue samples need to be processed. For one sample, 2 to 3 hr are sufficient for the first day’s work (Basic Protocol 1, steps 1 to 11). Medium changes can be completed in 20 min. The greatest time requirement in this protocol is for the chopping of the tissue samples. The hepatic differentiation experiment takes ∼1 hr on day 1 for coating the culture vessels. Counting the cells and seeding them on coated plates, the major task on day 2, can be completed in 1 hr. Medium change for differentiation experiments takes only several minutes. However, the media used for this experiment should be prepared and tested (for sterility and ability to support growth of cells) before the experiment is begun.

Literature Cited Chien, C.C., Yen, B.L., Lee, F.K., Lai, T.H., Chen, Y.C., Chan, S.H., and Huang, H.I. 2006. In vitro differentiation of human placenta-derived multipotent cells into hepatocyte-like cells. Stem Cells 24:1759-1768. Evarts, R.P., Nagy, P., Marsden, E., and Thorgeirsson, S.S. 1987. A precursor-product

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relationship exists between oval cells and hepatocytes in rat liver. Carcinogenesis 8:17371740.

Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.8.

In’t Anker, P.S., Scherjon, S.A., Kleijburg-van der Keur, C., de Groot-Swings, G.M., Claas, F.H., Fibbe, W.E., and Kanhai, H.H. 2004. Isolation of mesenchymal stem cells of fetal or maternal origin from human placenta. Stem Cells 22:13381345.

Schwartz, R.E., Reyes, M., Koodie, L., Jiang, Y., Blackstad, M., Lund, T., Lenvik, T., Johnson, S., Hu, W.S., and Verfaille, C.M. 2002. Multipotent adult progenitor cells from bone marrow differentiate into functional hepatocyte-like cells. J. Clin. Invest. 109:1291-1302.

Lee, K.D., Kuo, T.K., Whang-Peng, J., Chung, Y.F., Lin, C.T., Chou, S.H., Chen, J.R., Chen, Y.P., and Lee, O.K. 2004. In vitro hepatic differentiation of human mesenchymal stem cells. Hepatology 40:1275-1284.

Seo, M.J., Suh, S.Y., Bae, Y.C., and Jung, J.S. 2005. Differentiation of human adipose stromal cells into hepatic lineage in vitro and in vivo. Biochem. Biophys. Res. Commun. 328:258-264.

Kakinuma, S., Tanaka, Y., Chinzei, R., Watanabe, M., Shimizu-Saito, K., Teramoto, K., Arii, S., Sato, C., Takase, K., Yasumizu, T., and Teraoka, H. 2003. Human umbilical cord blood as a source of transplantable hepatic progenitor cells. Stem Cells 21:217-227.

Suzuki, A., Zheng, Y.W., Fukao, K., Nakauchi, H., and Taniguchi, H. 2004. Liver repopulation by c-Met positive stem/progenitor cells isolated from the developing rat liver. Hepatogastroenterology 51:423-426.

Kubota, H., Storms, R.W., and Reid, L.M. 2002. Variant forms of alpha-fetoprotein transcripts expressed in human hematopoietic progenitors: Implications for their developmental potential towards endoderm. J. Biol. Chem. 277:2762927635.

von Mach, M.A., Hengstler, J.G., Brulport, M., Eberhardt, M., Schormann, W., Hermes, M., Prawitt, D., Zabel, B., Grosche, J., Reichenbach, A., Muller, B., Weilemann, L.S., and Zulewski, H. 2004. In vitro cultured islet-derived progenitor cells of human origin express human albumin in severe combined immunodeficiency mouse liver in vivo. Stem Cells 22:1134-1141.

Petersen, B.E., Bowen, W.C., Patrene, K.D., Mars, W.M., Sullivan, A.K., Murase, N., Boggs, S.S., Greenberger, J.S., and Goff, J.P. 1999. Bone marrow as a potential source of hepatic oval cells. Science 284:1168-1170.

Yen, B.L., Huang, H.I., Chien, C.C., Jui, H.Y., Ko, B.S., Yao, M., Shun, C.T., Yen, M.L., Lee, M.C., and Chen, Y.C. 2005. Isolation of multipotent cells from human term placenta. Stem Cells 23:3-9.

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Isolation of Mesenchymal Stem Cells from Amniotic Fluid and Placenta

UNIT 1E.2

Shaun A. Steigman1 and Dario O. Fauza1 1

Department of Surgery, Children’s Hospital Boston and Harvard Medical School, Boston, Massachusetts

ABSTRACT Diverse progenitor cell populations, including mesenchymal, hematopoietic, trophoblastic, and possibly more primitive stem cells can be isolated from the amniotic fluid and the placenta. At least some of the amniotic and placental cells share a common origin, namely the inner cell mass of the morula. Indeed, most types of progenitor cells that can be isolated from these two sources share many characteristics. This unit will focus solely on the mesenchymal stem cells, the most abundant progenitor cell population found therein and, unlike some of the other stem cell types, present all through gestation. Protocols for isolation, expansion, freezing, and thawing of these cells are presented. Preference is given to the simplest methods available for any given procedure. Curr. C 2007 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 1:1E.2.1-1E.2.12.
Keywords: amniotic fluid r placenta r mesenchymal stem cells r fetus r neonate r amniotic stem cells r stem cells r tissue engineering r fetal tissue engineering

INTRODUCTION Both the amniotic fluid and the placenta contain a heterogeneous population of progenitor cells, which includes mesenchymal, hematopoietic, trophoblastic, and, perhaps, more primitive stem cells. At least some of the amniotic and placental cells share a common origin, namely the inner cell mass of the morula, which gives rise to the embryo, yolk sac, mesenchymal core of the chorionic villi, chorion, and amnion. Thus not surprisingly, most if not all types of progenitor cells that can be isolated from the amniotic fluid and the placenta share many characteristics. The cellular profile of the amniotic fluid changes predictably during gestation as the amniotic cavity receives cells from the fetus, and possibly from the placenta as well. The mechanisms responsible for the production and turnover of the amniotic fluid also contribute to the cell types present in the amniotic cavity. In addition to the spectrum of cells generally found in the amniotic fluid, certain fetal pathologic states (e.g., neural tube and body wall defects) may lead to the accessibility of cells not normally found therein. The multilineage potential of the different amniotic and placental stem cell populations has only recently begun to be explored. The most abundant population in amniotic fluid and nondecidual placenta, and the most extensively studied to date, is the mesenchymal stem cell (MSC), which can be isolated from these sources throughout gestation. Amniotic fluid and placental MSCs have been shown to differentiate at least into adipogenic, chondrogenic, myogenic, and osteogenic lineages. These cells can also give rise to nonmesenchymal cell types under appropriate conditions, raising the question as to whether “mesenchymal” is actually the best term to describe them. It is likely that their full differentiation spectrum remains to be entirely defined. The potential clinical value of amniotic fluid and placental MSCs in regenerative therapies has generated much interest of late. This unit will focus on MSCs only, as well as on the simplest methods available for any given procedure. Current Protocols in Stem Cell Biology 1E.2.1-1E.2.12 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e02s1 C 2007 John Wiley & Sons, Inc. Copyright


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The unit starts with the isolation and expansion of MSCs from either amniotic fluid (Basic Protocol 1) or placental samples (Basic Protocol 2). Cell isolation is achieved through a straightforward combination of natural selection by the culture medium and mechanical separation. Two means of implementing the mechanical separation are presented: multiple wells (Basic Protocols 1 and 2) or coverslips (Alternate Protocols 1 and 2). The unit proceeds to describe the basic methods for handling these MSCs, including passaging (Support Protocol 1) and freezing and thawing (Support Protocol 2), independent of whether they were isolated from placental or amniotic fluid sources. NOTE: Ethics approval for the described protocol is usually required from the appropriate institutional research office, typically an Institutional Review Board (IRB) for human cells or Institutional Animal Care and Use Committee (IACUC) for animal cells. NOTE: For all procedures described in this unit, sterile facilities for tissue culture and reagent preparation are required. If nondisposable instruments and containers are to be used, wash-up and sterilization facilities will also be required. NOTE: Experiments should be performed under sterile conditions in either Class II biological hazard flow hoods or laminar flow horizontal draft hoods. When working with human cells, Class II biological hazard flow hoods are recommended. NOTE: All incubations are performed in a humidified, 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

ISOLATION OF MESENCHYMAL STEM CELLS FROM AMNIOTIC FLUID Human amniotic fluid is typically obtained from diagnostic amniocentesis and occasionally from therapeutic amnioreductions. Conceivably, one could also procure it during delivery. Although the chemical and cellular composition of the amniotic fluid varies with gestational age, MSCs can be reliably isolated from it at any time during pregnancy. A very simple technique is outlined below for isolating amniotic fluid–derived MSCs (also referred to as mesenchymal amniocytes) based on natural selection from the culture medium and on mechanical separation.

Materials Amniotic fluid Mesenchymal-20 medium (see recipe) Dulbecco’s phosphate-buffered saline (cation-free; CMF-DPBS; Invitrogen) 0.025% (w/v) trypsin/0.04% (w/v) EDTA (Invitrogen) 15- and 50-ml conical centrifuge tubes (BD Biosciences) Centrifuge 6-well culture plate precoated with collagen type I (BD Biosciences) 10-cm tissue culture dishes (BD Biosciences) Inverted microscope Collect amniotic fluid sample 1. Place 40 ml amniotic fluid into 50-ml conical centrifuge tube. A sample size greater than 10 ml is recommended, particularly when initially implementing these protocols, although a minimal sample size of as little as 2 ml is feasible. For these small amounts of amniotic fluid, a 15-ml conical tube should be used instead.

2. Store amniotic fluid up to 48 hr at 4◦ C until ready to use. Isolation of Mesenchymal Stem Cells

Regardless of source, the amniotic fluid sample should be processed within 48 hr and preferably within 24 hr of harvest.

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Plate cells 3. Centrifuge the amniotic fluid sample 15 min at 500 × g, room temperature. 4. Aspirate the supernatant and discard. Resuspend the pellet in 6 ml mesenchymal-20 medium. 5. Plate 1 ml of the cell suspension into each well of a 6-well collagen-coated culture plate and incubate 48 hr. Upon inspection, mostly dead cells and cellular debris will be evident; typically, fewer than 2% of the cells therein are viable (see Fig. 1E.2.1). Instead of collagen, other options for coating the wells are fibronectin and laminin.

6. After 48 hr, change the medium by aspirating and replacing 1 ml per well. Mesenchymal amniocytes will adhere to collagen-coated plates by 48 hr; this step removes the nonadherent cells.

7. Aspirate and replace 1 ml of medium per well every 3 days thereafter.

Select MSCs 8. After 7 to 14 days, inspect individual wells for predominance of amniocytes with the characteristic “mesenchymal” morphology (see Fig. 1E.2.2) and for absence of contamination. Select only these wells for expansion. This time period can be fairly variable. Accordingly, daily plate inspections are recommended. The typical mesenchymal morphology is that of attached, spindle-shaped cells.

Figure 1E.2.1 Typical aspect of a fresh amniotic fluid sample after centrifugation, under trypan blue exclusion. Most cells are dead (blue cells); fewer than 2% of them are viable (unstained cells). Many cell debris are also present. (Magnification 100×)

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Figure 1E.2.2 Homogeneous population of amniotic fluid-derived mesenchymal cells after isolation, with the characteristic spindle-shaped morphology. (Magnification 100×)

Passage MSCs 9. When selected wells reach near confluence, aspirate medium and wash by adding 2 ml CMF-DPBS to each well and aspirating it. 10. Add 2 ml of 0.025% trypsin/0.04% EDTA, incubate 2 to 4 min to facilitate cell detachment, confirm detachment visually or under inverted microscope, and transfer each cell suspension into a separate 15-ml conical centrifuge tube. 11. Add 8 ml mesenchymal-20 medium to each tube. 12. Centrifuge each tube 5 min at 400 × g, room temperature. 13. Aspirate the supernatant carefully and resuspend the pellet in 6 ml mesenchymal-20 medium. 14. Dispense 1-ml aliquots of cell suspension into 10-cm tissue-culture dishes containing 9 ml mesenchymal-20 medium. 15. Grow cells to 80% to 90% confluence (replacing 9 to 10 ml of the medium every 3 days) and expand cultures (Support Protocol 1), or grow cells to ∼50% confluence and freeze (Support Protocol 2). ALTERNATE PROTOCOL 1

Isolation of Mesenchymal Stem Cells

ISOLATION OF MESENCHYMAL STEM CELLS FROM AMNIOTIC FLUID USING COVERSLIPS The mechanical separation component of the isolation protocol described above is achieved by choosing only those wells that contain predominantly the characteristic mesenchymal-appearing cells. An alternative means of mechanical separation is to use coverslips, instead of distinct wells. To that end, process the amniotic fluid as in Basic Protocol 1 through step 4. Then, distribute as many 5-mm2 coverslips (other sizes may also be used) as can fit on the surface of a single 10-cm collagen-coated plate and seed this plate at a cell density of 2 to 3 million cells per 150 cm2 (see Phelan, 2006). The coverslips can also be precoated with collagen. (As for the well-based method, other options for coating the culture plates and coverslips are fibronectin and laminin.) After 48 hr, inspect each individual coverslip, select those containing the morphologically distinct, mesenchymal-like cells, and place them into separate 30-cm2 culture plates containing

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6 ml fresh mesenchymal-20 medium. Replace the medium every 3 days and resume the remainder of Basic Protocol 1, beginning at step 9.

ISOLATION OF MESENCHYMAL STEM CELLS FROM PLACENTA Human placental tissue is typically procured from the whole organ, obtained at delivery, or from diagnostic chorionic villus sampling (CVS) specimens. Because the indications for CVS have progressively diminished recently and because of the sizeable variability of specimens produced by this method, the following text will refer to whole organ processing, i.e., from specimens obtained at delivery. Much of it can be adapted to the processing of CVS specimens, in accordance with the peculiarities of each individual CVS procurement. Once the placental tissue is cellularized (dissociated into a cell suspension), this protocol is identical to the one for amniotic fluid (Basic Protocol 1).

BASIC PROTOCOL 2

Materials Placental sample 10% type 2 collagenase (Worthington Biochemical) Dispase II (Roche Applied Science) CaCl2 Mesenchymal-20 medium (see recipe) Dulbecco’s phosphate-buffered saline (cation-free; CMF-DPBS; GIBCO) 0.025% (w/v) trypsin/0.04% (w/v) EDTA (Invitrogen) Serum-free DMEM (Sigma) 15- or 50-ml conical centrifuge tubes (BD Biosciences) 100-µm mesh (Fisher Scientific) Centrifuge 6-well culture plate precoated with collagen type I (BD Biosciences) Inverted microscope 10-cm tissue culture dishes (BD Biosciences) Process placental sample 1. Mechanically remove the maternal decidua by peeling it away from the remainder of the placenta, which contains the chorionic villi (each with a mesenchymal core), and discard. 2. Mince the residual specimen (chorionic villi with mesenchymal core) and transfer it to a 15-or 50-ml conical tube (depending on the volume of the sample). 3. Add a solution containing 10% (w/v) type 2 collagenase and 4.0 U dispase II/2.5 mmol CaCl2 per liter to cover the tissue. The optimal amount of enzyme mixture per weight of tissue has not yet been established.

4. Filter the mixture through 100-µm mesh into a 15-ml conical centrifuge tube. 5. Centrifuge 15 min at 500 × g, room temperature. 6. Resuspend the pellet in 10 ml mesenchymal-20 medium. The placental samples that the authors have processed ranged in weight between 12 and 19 g, and the resulting pellet could be resuspended in 10 ml medium. The optimal volume of medium per weight of tissue has not yet been established.

Isolate MSCs 7. Plate 1 ml of the cell suspension into each well of a 6-well, collagen-coated culture plate, and place in incubator. Instead of collagen, other options for coating the wells are fibronectin and laminin.

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8. After 48 hr, change the medium by aspirating and replacing 1 ml per well MSCs will adhere to collagen-coated plates by 48 hr; this step removes the nonadherent cells.

9. Aspirate and replace 1 ml of medium per well every 3 days thereafter.

Select MSCs 10. After 7 to 14 days, inspect individual wells for predominance of cells with the characteristic mesenchymal morphology (see Fig. 1E.2.2) and for absence of contamination. Select only these wells for expansion. This time period can be fairly variable. Accordingly, daily plate inspections are recommended. The typical mesenchymal morphology is of attached, spindle-shaped cells.

Passage cells 11. When selected wells reach near confluence, aspirate medium, add 2 ml CMF-DPBS to each well, and aspirate the buffer. 12. Add 2 ml of 0.025% trypsin/0.04% EDTA and incubate 2 to 4 min to facilitate cell detachment. 13. Confirm detachment visually or under an inverted microscope and transfer each cell suspension into separate 15-ml conical centrifuge tubes. 14. Add 8 ml mesenchymal-20 medium to each tube. 15. Centrifuge each tube 5 min at 400 × g, room temperature. 16. Aspirate the supernatant carefully and discard. Resuspend the pellet in 6 ml mesenchymal-20 medium. 17. Add 1 ml cell suspension to each of 10-cm tissue-culture dishes containing 9 ml of mesenchymal-20 medium. 18. Grow cells to 80% to 90% confluence (replacing 9 to 10 ml of the medium every 3 days) and expand cultures (Support Protocol 1), or grow cells to ∼50% confluence and freeze (Support Protocol 2). ALTERNATE PROTOCOL 2

ISOLATION OF MESENCHYMAL STEM CELLS FROM PLACENTA USING COVERSLIPS The mechanical separation component of the isolation protocol described above is achieved by choosing only those wells that contain predominantly the characteristic mesenchymal-appearing cells. An alternative means of mechanical separation is to use coverslips, instead of distinct wells. To that end, process the placental tissue as in Basic Protocol 2 through step 5. Then, distribute as many (usually, but not necessarily, 5 mm2 ) coverslips as can be fit on the surface of a single 10-cm collagen-coated plate and seed this plate at a cell density of 2 to 3 million cells per 150 cm2 (see Phelan, 2006). The coverslips can also be precoated with collagen. (As for the well-based method, other options for coating the culture plates and coverslips are fibronectin and laminin.) After 48 hr, inspect each individual coverslip, select those containing the morphologically distinct, mesenchymal-like cells, and place them into separate 30-cm2 culture plates containing fresh 6 ml mesenchymal-20 medium (see recipe). Change the medium every 3 days and resume the remainder of Basic Protocol 2, beginning at step 11.

Isolation of Mesenchymal Stem Cells

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EXPANSION OF MESENCHYMAL STEM CELLS FROM AMNIOTIC FLUID OR PLACENTA

SUPPORT PROTOCOL 1

After isolation, cells will typically reach confluence in 3 to 5 days and require passage. For passages beyond p2, the authors typically lower the serum content of the medium from 20% to 10%, solely for cost savings because these cells expand quite rapidly even at 10%. This protocol describes the technique for passaging cells that have achieved 80% to 90% confluence.

Materials MSC in culture at 80% to 90% confluence Dulbecco’s phosphate-buffered saline (cation-free; CMF-DPBS; Invitrogen) 0.025% (w/v) trypsin/0.04% (w/v) EDTA (Invitrogen) Mesenchymal-10 medium (see recipe) Inverted microscope 10-cm tissue culture dishes (BD Biosciences) 1. Aspirate medium from cultured cells, using a Pasteur pipet. 2. Wash the plate with 2 ml of either CMF-DPBS or 0.025% trypsin/0.04% EDTA: apply the solution gently along side of dish, swirl, and immediately aspirate and discard. 3. Add 3 ml 0.025% trypsin/0.04% EDTA and return the plate to the incubator for ∼2 to 4 min to allow detachment of the cells. 4. Confirming detachment by observing under an inverted microscope. 5a. If splitting cells 2:1: Load 10-cm tissue culture dishes with 6 ml mesenchymal-10 medium. Add 5 ml mesenchymal-10 medium to the trypsinized plate, mix with gentle repeated pipetting for ∼30 sec, and aliquot 4 ml into each of the two prepared dishes. 5b. If splitting cells 3:1: Load 10-cm tissue culture dishes with 7 ml mesenchymal-10 medium. Add 6 ml mesenchymal-10 medium to the trypsinized plate, mix with gentle repeated pipetting for ∼30 sec, and dispense 3 ml into each of the three prepared dishes. 5c. If splitting cells 4:1: Load 10-cm tissue culture dishes with 8 ml mesenchymal-10 medium. Add 5 ml mesenchymal-10 medium to the trypsinized plate, mix with gentle repeated pipetting for ∼30 sec, and dispense 2 ml into each of the four prepared dishes. 6. Return dishes to the incubator. Grow cells to 80% to 90% confluence (replacing 9 to 10 ml of the medium every 3 days) and passage by repeating the steps in this protocol, or grow cells to ∼50% confluence and freeze (Support Protocol 2).

FREEZING AND THAWING MESENCHYMAL STEM CELLS From early passage through any point during MSC expansion, these cells can be frozen for future use. The time span for their viability at −80◦ C has yet to be reliably determined, but it is certainly possible to store them for at least a year. Freezing of MSCs is optimally performed with exponentially growing cells, typically near 50% confluence.

SUPPORT PROTOCOL 2

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Materials MSC in culture, ∼50% confluent Dulbecco’s phosphate-buffered saline (cation-free; CMF-DPBS; Invitrogen) 0.025% (w/v) trypsin/0.04% (w/v) EDTA (Invitrogen) Mesenchymal-10 medium (see recipe) Freezing solution: add 2 ml DMSO to 8 ml mesenchymal-10 medium DMSO (Sigma-Aldrich) Pasteur pipet Inverted microscope 15-ml conical centrifuge tube (BD Biosciences) Centrifuge 2.0 ml cryogenic vials (Fisher Scientific) –80◦ C freezer 37◦ C water bath 10-cm tissue culture dishes (BD Biosciences) Freeze MSCs 1. Aspirate medium from the 50% confluent MSC culture, using a Pasteur pipet. 2. Wash the plate with 2 ml of either CMF-DPBS or 0.025% trypsin/0.04% EDTA: apply the solution gently along the side of dish, swirl, and immediately aspirate. 3. Add 3 ml 0.025% trypsin/0.04% EDTA and return the plate to the incubator for ∼2 to 4 min to allow detachment of the cells. 4. Confirm detachment by observing under an inverted microscope. 5. Add 7 ml mesenchymal-10 medium to neutralize trypsin, bringing total volume to 10 ml, mix with gentle repeated pipetting for ∼30 sec, and transfer the cell suspension to a 15-ml conical centrifuge tube. 6. Centrifuge 5 min at 200 × g, room temperature. 7. Aspirate the supernatant and discard. Resuspend the pellet in 1.5 ml mesenchymal-10 medium. 8. Add 1.5 ml freezing solution and mix with gentle repeated pipetting for ∼30 sec. CAUTION: DMSO is hazardous. When preparing the freezing solution, work in a fume hood and use gloves.

9. Dispense 1-ml aliquots into each of three cryogenic vials. 10. Store up to 1 year at −80◦ C.

Thaw MSCs 11. Remove vial from the −80◦ C freezer and roll the vial between the hands for 10 to 15 sec until the outside of vial is frost free. 12. Hold the cryogenic vial in 37◦ C water bath until the contents are visibly thawed. To protect sterility do not submerge the cap.

13. Immerse the vial in 95% ethanol bath or spray with 70% ethanol to kill microorganisms from water bath and air dry in sterile hood. 14. Dispense the contents into a 10-cm tissue culture plate containing 9 ml mesenchymal10 medium. Isolation of Mesenchymal Stem Cells

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Fetal bovine serum (FBS) stock Divide samples of fetal bovine serum, MSC-qualified (Invitrogen) into 50 ml aliquots and store at −20◦ C. When ready for use, thaw a 50-ml aliquot of FBS in 37◦ C water bath. Optional: Prior to adding to media, sterilize by passing through 0.22-µm polyethersulfone sterilizing, low protein–binding membrane (Corning), using a vacuumdriven filter system. FBS purchased from Hyclone has traditionally been used in the authors’ laboratory for MSC cultures, but recently it has been observed that MSC-qualified serum from Invitrogen leads to more robust cell growth. Batch testing of all serum is imperative to ensure optimal expansion of MSCs. Avoid multiple freeze-thaw cycles.

Mesenchymal-10 and mesenchymal-20 medium 500 ml high-glucose DMEM containing L-glutamine (Sigma) supplemented with: 10% or 20% (v/v) FBS stock (see recipe) 5 ml 10,000 U/ml penicillin G sodium/10 mg/ml streptomycin sulfate antibiotic solution (Sigma; final concentration 100 U/ml PCN G and 0.1 mg/ml streptomycin) 50 µl 50 µg/ml rhFGF-basic stock (see recipe; final concentration 5 ng/ml) Store up to 3 weeks at 4◦ C The recipe name reflects the FBS concentration in the medium (10% or 20%).

rhFGF-basic stock, 50 µg/ml Reconstitute 25 µg recombinant human, basic, fibroblast growth factor (Promega) in 500 µl sterile water. Divide into 50-µl aliquots and store up to 1 year at −20◦ C. Avoid multiple freeze-thaw cycles.

COMMENTARY Background Information Ever since Friedenstein’s original description of colony-forming, spindle-shaped stromal cells in the bone marrow with multipotent differentiation potential, MSCs have been intensively studied for over 40 years (Friedenstein et al., 1966, 1968, 1970). The plasticity, self-renewal, and multilineage potentials of MSCs (Prockop, 1997; Pittenger et al., 1999) have generated increasing interest in their use in an ever expanding variety of regenerative therapy applications. At the same time, a plethora of reports have identified MSCs in diverse sources (Vaananen, 2005). Most of these sources, however, would not be compatible with perinatal cell-based therapies. In the perinatal setting, ethical objections to the isolation of amniotic fluid and

placental MSCs should be obviated by the fact that amniocentesis and chorionic villus sampling are widely performed diagnostic procedures. An amniocentesis is the safest of any invasive prenatal diagnostic method, being associated with a less than 0.5% spontaneous abortion rate (Jauniaux and Rodeck, 1995). Furthermore, a mother carrying a fetus with a congenital anomaly on prenatal imaging is routinely offered a diagnostic amniocentesis. In this instance, a small extra aliquot of amniotic fluid could be obtained at that time for tissue engineering, gene therapy, or cell transplantation purposes without any additional risk to the mother or fetus. Along these lines, the authors of this unit have recently introduced, and continued to develop, the concept of using MSCs derived from

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amniotic fluid for tissue engineering strategies for the surgical repair of congenital anomalies in the perinatal period. Translated into the clinical context, from a simple amniocentesis grafts could be engineered in parallel to the remainder of gestation so that a newborn, or even a fetus, could benefit from having autologous tissue promptly available for surgical reconstruction, either before or at birth. Thus far, the authors have validated this concept in large animal models of diaphragmatic and tracheal repairs (Fuchs et al., 2004; Kunisaki et al., 2006a,b). Cells from all three germ layers have been identified in the amniotic fluid for nearly 30 years (Milunsky, 1979; Hoehn and Salk, 1982; Gosden, 1983; Prusa et al., 2003). However, the isolation of progenitor cells was first reported only in 1993, when small, nucleated, round cells identified as hematopoietic progenitor cells were found therein before week 12 of gestation (Torricelli et al., 1993). The multilineage potential of nonhematopoietic stem cells was first suggested in 1996 by the demonstration of myogenic conversion of amniocytes (Streubel et al., 1996). Nonetheless, that study did not specify the identity of the cells that responded to the myogenic culture conditions, namely the supernatant of a rhabdomyosarcoma cell line. The proper characterization, differentiation potential, and therapeutic applications of mesenchymal amniocytes have started to be determined only quite recently (Kaviani et al., 2001; 2003; Int ‘t Anker et al., 2003; Fuchs et al., 2004; Kunisaki et al., 2006a,b,c). Because of the mechanisms behind placental development, different cell types are found there at different gestational ages (Fauza, 2004). At approximately week 5 postconception, all placental villi are of the mesenchymal type. Much like mesenchymal amniocytes, placental mesenchymal cells have started to be well identified and explored only of late (Haigh et al., 1999; Kaviani et al., 2002). In addition to the deliberately simple and easily reproducible methodology for isolation of amniotic fluid and placental MSCs described here, other methods that rely on oxygen tension, two-stage cultures, or alternative media formulations have been reported (Tsai et al., 2004; In ‘t Anker et al., 2004). Also, it remains to be determined whether amniotic stem cells recently obtained by somewhat different methods and described as more primitive than purely mesenchymal are actually the same cells commonly referred to as amniotic

MSCs, which have been shown to be able to give rise to cells from more than one germ layer (Tsai et al., 2006; De Coppi et al., 2007). Finally, more recently, as a possible prerequisite to eventual clinical trials of amniotic MSC-based therapies, the authors have recently isolated and characterized human MSCs in the absence of fetal bovine serum (Kunisaki et al., 2007).

Critical Parameters and Troubleshooting All experiments should be performed under sterile conditions in either Class II biological hazard flow hoods or laminar flow horizontal draft hoods. When working with human cells, Class II biological hazard flow hoods are recommended. Appropriate sterilization measures for reagents and containers are a necessity. Although amniotic MSCs, on the spectrum of cell culture, are fairly hardy cell lines, the author’s experience has shown that strict adherence to sterile techniques are a must. Batch testing of all serum is also a requirement to validate the purity of the batch and its ability to maintain undifferentiated MSCs in culture (characterized by their spindle shape and presence or absence of molecular markers; see below). Upon delivery, FBS should be tested, divided into aliquots, and stored at −20◦ C. In this manner, FBS should only be thawed once because multiple freeze/thaw cycles impede its efficacy for cell culture. Media can typically be stored up to 2 or 3 weeks and should be inspected prior to each use. The color of the medium, as a reflection of pH, will evolve from pink-amber (fresh) to pink-magenta (old). In addition, evidence of crystallization or flocculation should prompt discarding. When first performing MSCs isolation, it is prudent to confirm MSC identity. Such steps in stem cell characterization are outside the scope of this particular unit on isolation, but MSCs display a typical pattern of cell surface markers compatible with a multipotent mesenchymal progenitor lineage. The markers include CD73 (SH3), CD105 (SH2), CD44, CD29, CD90, CD13, CD10, and CD71. MSCs are also positive for human leukocyte antigens A, B, and C and negative for CD45, CD34, CD14, CD19, CD8, CD56, and CD31 (see Pittinger et al., 1999; Kaviani et al., 2001, 2003; Kunisaki et al., 2007).

Anticipated Results MSC isolation via the protocols in this unit is fairly straightforward and consistently

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reliable. However, as stated in the introduction, the cellular and chemical context of the amniotic fluid varies with gestational age, which may lead to variations in initial cell density between samples. Thus, there can be no “expected” or “typical” yield of MSCs from the initial isolation to be used as a guide. Regardless, the initial yield is not the ratelimiting step because amniotic fluid samples procured during pregnancy can generate sufficient numbers of MSCs within just a few generations. Once derived, MSCs have lengthy lifespans and have been carried in the authors’ laboratory for up to 45 passages. Nonetheless, siphoning off early-passage MSCs for freezing is routinely performed.

Time Considerations The simplicity of these isolation protocols is among their key advantages. Overall time commitment for each step is not overburdening. Typically, it takes 2 to 3 weeks to generate ample amounts of MSCs from a small sample of amniotic fluid or placenta. The amounts of FBS and FGF added to the culture media have a direct impact on cell kinetics. Placental dissection The mechanical separation of the decidual layer from the remainder of the placental specimen, which contains the chorionic villi, generally takes about 1 hr but will depend on the familiarity of the operator with basic tissue dissection technique and placental anatomy (see Internet Resources). Dissolving, filtering, and resuspending the tissue into single cells can take up to another 1 hr. Initial MSC isolation The initial seeding of the amniotic or placental sample onto the precoated plates takes ∼45 min. MSC expansion The transfer of selected MSCs from the precoated plates to the tissue culture dishes requires ∼45 min. Media preparation Preparation of media typically takes 30 min and needs to be performed every 2 to 3 weeks. Working with MSCs Passaging and freezing each require ∼30 min, depending on the number of plates being processed. Thawing requires 5 min, beyond the setup time for cleaning the workspace and bringing reagents to room temperature.

Literature Cited De Coppi, P., Bartsch Jr., G., Siddiqui, M.M., Xu, T., Santos, C.C., Perin, L., Mostoslavsky, G., Serre, A.C., Snyder, E.Y., Yoo, J.J., Furth, M.E., Soker, S., and Atala, A. 2007. Isolation of amniotic stem cell lines with potential for therapy. Nat. Biotechnol. 25:100-106. Fauza, D.O. 2004. Amniotic fluid and placental stem cells. In Best Practice & Research Clinical Obstetrics & Gynaecology, Vol. 18: Stem Cells in Obstetrics and Gynaecology (N.M. Fisk and J. Itskovitz-Eldor, eds.) pp. 877-891. Elsevier, London. Friedenstein, A.J., Piatetzky-Shapiro I.I., and Petrakova, K.V. 1966. Osteogenesis in transplants of bone marrow cells. J. Embryol. Exp. Morphol. 3:381-390. Friedenstein, A.J., Petrakova, K.V., Kurolesova, A.I., and Frolova, G.P. 1968. Heterotopic bone marrow. Analysis of precursor cells for osteogenic and hematopoietic tissues. Transplantation 2:230-247. Friedenstein, A.J., Chailakhgan, R.K., and Lalykina, K.S. 1970. The development of fibroblast colonies in monolayer cultures of guinea pig bone marrow and spleen cells. Cell Tissue Kinet. 20:263-272. Fuchs, J.R., Kaviani, A., Oh, J.T., LaVan, D., Udagawa, T., Jennings, R.W., Wilson, J.M., and Fauza, D.O. 2004. Diaphragmatic reconstruction with autologous tendon engineered from mesenchymal amniocytes. J. Pediatr. Surg. 39:834-838. Gosden, C.M. 1983. Amniotic fluid cell types and culture. Br. Med. Bull. 39:348-354. Haigh, T., Chen, C., Jones J., and Aplin, J. D. 1999. Studies of mesenchymal cells from 1st trimester human placenta: Expression of cytokeratin outside the trophoblast lineage. Placenta 20:615625. Hoehn, H. and Salk, D. 1982. Morphological and biochemical heterogeneity of amniotic fluid cells in culture. Methods Cell Biol. 26:11-34. Int ‘t Anker, P.S., Scherjon, S.A., Kleijburg-van der Keur, C., Noort, W.A., Claas, F.H.J., Wilemze, R., Febbe, W.E., and Kanhai, H.H.H. 2003. Amniotic fluid as a novel source of mesenchymal stem cells for therapeutic transplantation. Blood 102:1548-1549. Int ‘t Anker, P.S., Scherjon, S.A., Kleijburg-van der Keur, C., de Groot-Swings, G.M.J.S., Claas, F.H.J., Wilemze, R., Febbe, W.E., and Kanhai, H.H.H. 2004. Isolation of mesenchymal stem cells of fetal or maternal origin from human placenta. Stem Cells 22:1338-1345. Jauniaux, E. and Rodeck, C. 1995. Use, risks, and complications of amniocentesis and chorionic villus samples for prenatal diagnosis in early pregnancy. Early Pregnancy 1:245-252. Kaviani, A., Perry, T.E., Dzakovic, A., Jennings, R.W., Ziegler, M.M., and Fauza, D.O. 2001. The amniotic fluid as a source of cells for fetal tissue engineering. J. Pediatr. Surg. 36:1662-1665.

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Kaviani, A., Perry, T.E., Barnes, C.M., Oh, J.T., Zeigler, M.M., Fishman, S.J., and Fauza, D.O. 2002. The placenta as a cell source in fetal tissue engineering. J. Pediatr. Surg. 37:995-999. Kaviani, A., Guleserian, K., Perry, T.E., Jennings, R.W., Zeigler, M.M., and Fauza, D.O. 2003. Fetal tissue engineering from amniotic fluid. J. Am. Coll. Surg. 196:592-597. Kunisaki, S.M., Freedman, D.A., and Fauza, D.O. 2006a. Fetal tracheal reconstruction with cartilaginous grafts engineered from mesenchymal amniocytes. J. Pediatr. Surg. 41:675682. Kunisaki, S.M., Fuchs, J., Kaviani, A., Oh, J.T., LaVan, D., Vacanti, J.P., Wilson, J.M., and Fauza, D.O. 2006b. Diaphragmatic repair through fetal tissue engineering: A comparison between mesenchymal amniocyte- and myoblast-based constructs. J. Pediatr. Surg. 41:34-39. Kunisaki, S.M., Jennings R.W., and Fauza D.O. 2006c. Fetal cartilage engineering from amniotic mesenchymal progenitor cells. Stem Cells Dev. 15:245-253. Kunisaki, S.M., Armant, M., Kao, G.C., Stevenson, K., Kim, H., and Fauza, D.O. 2007. Tissue engineering from human mesenchymal amniocytes: A prelude to clinical trials. J. Pediatr. Surg. In press. Milunsky, A. 1979. Amniotic fluid cell culture. In Genetic Disorders of the Fetus (A. Milunsky, ed.) pp. 75-84. Plenum Press, New York. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.8. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., and Marshak, D.R. 1999. Multilineage potential of adult hu-

man mesenchymal stem cells. Science 284:143147. Prockop, D.J. 1997. Marrow stromal cells as stem cells for nonhematopoietic tissues. Science 276:71-74. Prusa, A.R., Marton, E., Rosner, M., Bernaschek, G., and Hengtschlager, M. 2003. Oct-4expressing cells in human amniotic fluid: A new source for stem cell research? Hem. Reprod. 18:1489-1493. Streubel, B., Martucci-Ivessa, F., Fleck, T., and Bittner R.E. 1996. In vitro transformation of amniotic cells to muscle cells–background and outlook. Wien Med. Wochenschr. 146:216217. Torricelli, F., Brizzi, L., Bernabei, P.A., Gheri, G., Di Lollo, S., Nutini, L., Lisi, E., Di Tomasso, M., and Cariati, E. 1993. Identification of hematopoietic progenitor cells in human amniotic fluid before the 12th week of gestation. Ital. J. Anat. Embryol. 98:119-126. Tsai, M.S., Lee, J.L., Chang, Y.J., and Hwang, S.M. 2004. Isolation of human multipotent mesenchymal stem cells from second-trimester amniotic fluid using a novel two-stage culture protocol. Hum. Reprod. 19:1450-1456. Tsai, M.S., Hwang, S.M., Tsai, Y.L., Cheng, F.C., Lee, J.L., and Chang, Y.J. 2006. Clonal amniotic fluid-derived stem cells express characteristics of both mesenchymal and neural stem cells. Biol. Reprod. 74:545-51. Vaananen, H.K. 2005. Mesenchymal stem cells. Ann. Med. 37:469-479.

Internet Resources http://www.simba.rdg.ac.uk/Dave/Lit%20review. html This Web site contains a useful drawing of placental structure.

Isolation of Mesenchymal Stem Cells

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Isolation of Amniotic Epithelial Stem Cells

UNIT 1E.3

Toshio Miki,1 Fabio Marongiu,1 Kenneth Dorko,1 Ewa C.S. Ellis,1 and Stephen C. Strom1 1

University of Pittsburgh, Pittsburgh, Pennsylvania

ABSTRACT Many of the cell types that can be isolated from placental tissues retain phenotypic plasticity that makes them an interesting source of cells for regenerative medicine. Several procedures for the isolation of stem cells from different parts of the placenta have been reported. This unit describes a detailed and simple protocol for the selective isolation of amniotic epithelial cells from human term placenta without disturbing the mesenchymal layer. We also introduce a simple density separation technique for the enrichment of the population for SSEA-4 positive cells. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 12:1E.3.1-1E.3.10.  Keywords: placenta r amnion r epithelial stem cells

INTRODUCTION The amnion is a thin, avascular membrane composed of both cuboidal and columnar epithelial cells, which are in contact with the amniotic fluid on the external side and attached to a basal lamina on the inner side. This lamina is connected to the amniotic mesoderm; it is a layer of extracellular components (collagens and fibronectin) in which are embedded a network of dispersed fibroblast-like mesenchymal cells (Fig. 1E.3.1). Based on immunohistochemical and genetic analysis, human amniotic epithelial cells (hAECs) have the potential to differentiate into all three germ layers: endoderm (liver, pancreas), mesoderm (cardiomyocyte), and ectoderm (neural cells) in vitro. The availability of hAECs and the absence of ethical concerns for this source of stem cells are considered advantageous for their widespread use and acceptance. Amnion derived from term placenta following live birth may be a useful and noncontroversial source of stem cells for cell transplantation and regenerative medicine. In this unit, we describe the method to selectively isolate human amniotic epithelial cells from term placenta obtained from cesarean section procedures (Basic Protocol). Furthermore, we introduce a method to enrich the population for SSEA-4 positive cells to >97% by a simple density separation technique using Percoll (Alternate Protocol). NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: This protocol uses human tissues. It should be submitted to, reviewed by, and approved by the appropriate Institutional Review Board, and all tissues should be obtained with the informed consent of the source.

ISOLATION OF HUMAN AMNIOTIC EPITHELIAL CELLS Human amniotic epithelial cells (hAECs) are isolated from term placentas which would normally be discarded after delivery. For sterility purposes, placentas are normally obtained from caesarean section; however, theoretically, any placenta should be useful for Current Protocols in Stem Cell Biology 1E.3.1-1E.3.10 Published online January 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e03s12 C 2010 John Wiley & Sons, Inc. Copyright 

BASIC PROTOCOL Embryonic and Extraembryonic Stem Cells

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epithelial cells

mesenchymal cells

Figure 1E.3.1

Cross-section of human amnion.

amniotic epithelial cell isolation. Tissues are obtained with local Institutional Review Board (IRB) approval in the U.S., or under appropriate Ethical Committee approval. For the studies described here, all infectious pathogen–positive deliveries including those involving HBV, HCV, and HIV, as well as cases of prediagnosed genetic abnormalities, are excluded. Even with these precautions, all staff members must be made aware that the tissue should be considered potentially infectious material and standard precautions for safe use of human tissue must be followed.

Materials Hanks’ balanced salt solution (HBSS; calcium- and magnesium-free; CMF-HBSS; Lonza, cat. no. 04-315Q), sterile Term placenta, freshly delivered, in sterile transportation medium (see recipe) 70% ethanol 0.05% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25300-054) Pre-digestion buffer (see recipe), sterile Standard culture medium (see recipe), sterile

Isolation of Amniotic Epithelial Stem Cells

Laminar flow cabinet (BSL-2) equipped with the following: Absorbent bench paper Sterile field, 16 × 29 in. Sterile scalpel 500-ml glass beakers (4) Sterile scissors (2) and forceps (2), Sterile gloves and sleeves 100-μm nylon cell strainers (4) 50-ml conical polypropylene (e.g., Falcon) centrifuge tubes (8) Refrigerated centrifuge Additional reagents and equipment for cell counting (UNIT 1C.3)

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Figure 1E.3.2

Sterile equipment for the isolation of hAECs.

Prepare hood for isolation 1. Equip a BSL-2 laminar-flow cabinet with the following (Fig. 1E.3.2): Absorbent bench paper Sterile field, 16 × 29 inches Sterile scalpel Sterile 500-ml beaker Sterile scissors and forceps Sterile gloves and sleeves 100-μm nylon cell strainers, sterile 50-ml Falcon tubes, sterile. 2. Add 200 ml CMF-HBSS to a 500-ml beaker.

Carry out dissection 3. Wearing sterile gloves, place the whole placenta on the sterile field (Fig. 1E.3.3). The maternal surface (rough surface) should be facing down on the paper, with the smooth surface bearing the umbilical cord facing up. In this position, the amnion membrane will lay across the upper surface of the placenta.

4. Trim the umbilical cord close to the placental surface and cut an X-shaped incision into, but not through, the placental tissue (Fig. 1E.3.4). The X should intersect at the position of the umbilical cord. It may be easiest to start the incision in the region of the umbilical cord. After this step there could be an excess of blood on top of the amnion. If that is the case, pour some CMF-HBSS on top of the membrane and gently massage to remove blood clots. Embryonic and Extraembryonic Stem Cells

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Figure 1E.3.3

Placenta with amnion membrane facing up.

Figure 1E.3.4

Placenta with an X-shaped incision.

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Figure 1E.3.5

Peeling the amnion membrane.

5. Peel the amnion membrane from the underlying chorion layer of the placenta body. Start from the cut edge (middle of the placenta body) and peel the membrane from the placenta (Fig. 1E.3.5). While peeling the amnion, it should look like a matte see-through membrane. Should the surface look rough, you are peeling off some extra connective tissue at the interface between amnion and chorion. This will likely interfere with the isolation, resulting in low yield and inefficient centrifugation. In order to avoid taking extra tissue, pinch the rough part with a second pair of forceps and only collect the amnion. This phenomenon is more common in tissue from lengthy labor or when the amnion is partially detached from the discoid placenta.

6. Place the amnion in the sterilized 500-ml glass beaker containing 200 ml CMFHBSS, from step 2 (Fig. 1E.3.6). 7. Discard the remaining part of the placenta, remove the dissecting material from the work area, and wipe the hood with 70% ethanol. 8. Wash the amnion two to three times, each time by moving the amnion to a clean 500-ml beaker containing ∼200 ml fresh CMF-HBSS. This washing step is crucial for the trypsin to work properly. Blood clots will reduce the efficiency of the trypsin. Ideally, all clots on the membrane will be cleared.

Digest the membrane 9. Thaw and prewarm the trypsin/EDTA solution to 37◦ C in a water bath. 10. Place the membrane in a 50-ml centrifuge tube and add 20 ml pre-digestion buffer. Gently rock the membrane in the solution for 30 sec, then transfer the membrane with forceps to two new 50-ml tubes. Discard the buffer. The volume of membrane tissue from an average placenta is 20 ml.

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Figure 1E.3.6

Washing the amniotic membrane in HBSS.

11. Add 20 ml of pre-digestion buffer to the tissue in the tubes. Incubate 10 min at 37◦ C. 12. Transfer the membrane into new tubes (discarding the pre-digestion buffer) and add 30 ml trypsin/EDTA solution. Incubate for 40 min at 37◦ C. 13. Transfer the membrane into new 50-ml tubes, add ∼30 ml fresh trypsin/EDTA, and incubate for an additional 40 min. Save the trypsin digest. 14. Add two volumes of standard culture medium to the trypsin digest from step 12 and centrifuge 10 min at ∼200 × g, 4◦ C. 15. Decant the supernatant and resuspend the pellet in 10 ml of standard culture medium. The cell pellet might appear like a compact tissue-like aggregate. Pipet up and down several times in order to release the cells. Discard any remaining small cell aggregates or filter the cell preparation through a 100-μm filter. If necessary, centrifuge one or two more times at ∼200 × g, 4◦ C.

16. Perform steps 14 and 15 again when the second 40-min digest (from step 13) is complete, then pool the two digests in one 50-ml tube.

Prepare cultures 17. Count the cell number (UNIT 1C.3). 18. If cells are to be cultured, plate up to 1 × 105 cells per cm2 in standard culture medium. At this point, cells can be used for flow cytometric analysis, plated for in vitro experimentation, or further enriched for SSEA 4–expressing cells (Alternate Protocol). Isolation of Amniotic Epithelial Stem Cells

For our experiments, amniotic epithelial cells are initially cultured in the standard medium described in the Reagents and Solutions section, containing DMEM medium, or a version of this medium formulated for knock-out experiments (K-DMEM; also see recipe

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in Reagents and Solutions). We do not know that these are the best media for all applications, so each laboratory should determine empirically the best culture conditions for the differentiated state that they wish to maintain. In DMEM medium supplemented as described in Reagents and Solutions, with 10% serum and 10 ng/ml epidermal growth factor (EGF), the cells grow for 2 to 6 passages, a passage being approximately a 1:4 split when the cells approach confluence. Removal of the EGF results in a rapid decrease in proliferation and what appears to be terminal differentiation. Even in the continued presence of serum and EGF, the cells do not normally proliferate past passage 6 under the culture conditions described here. Although not optimized at this time, like embryonic stem cells (ESC), hAECs can be induced to differentiate to specific cell types including neurons (Sakuragawa et al., 1996, 1997; Kakishita et al., 2000, 2003; Elwan et al., 2003a,b; Elwan and Sakuragawa, 2007), cardiomyocytes (Miki et al., 2005), pancreatic cell types (Wei et al., 2003; Miki et al., 2005), and hepatocytes (Takashima et al., 2004; Miki et al., 2005). Culture conditions used to induce differentiation along these pathways are in the original publications.

ENRICHMENT OF STEM CELL MARKER–POSITIVE CELLS BY DENSITY SEPARATION

ALTERNATE PROTOCOL

One of the common embryonic stem cell surface markers is a stage-specific embryonic antigen, SSEA-4, the antibody for which was raised against a human teratocarcinoma cell line. The expression of SSEA-4 is normally down-regulated when stem cells differentiate. In this section, we describe a density separation method to enrich for SSEA-4 positive cells from human amniotic epithelial cells.

Additional Materials (also see Basic Protocol) Isolated amniotic epithelial cells (Basic Protocol, steps 1 to 16) 24% (v/v) Percoll solution (see recipe), sterile 1. Suspend isolated amniotic epithelial cells (Basic Protocol, step 16) in CMF-HBSS at a density of 5 × 106 /ml. 2. Gently pipet 20 ml of the cell suspension onto a cushion of 24 ml of 24% Percoll in a 50-ml conical tube. 3. Centrifuge 8 min at ∼150 × g, 21◦ C. Stop without using the brake. 4. Carefully discard the supernatant without disturbing the cell pellet. If the pellet is too small and the middle band is too thick, aspirate and transfer the band into a new tube. Repeat this protocol in order to recover more cells.

Figure 1E.3.7

Density separation for stem cell enrichment.

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5. Resuspend the pellet in CMF-HBSS (up to 50 ml), mix, centrifuge 5 min at ∼150 × g, 21◦ C, and remove the supernatant, to wash the cells. Most of the dead and SSEA-4 negative cells are separated out by this procedure and the total cell number will decrease to 30% to 40% of the original cell number. The results of a typical separation are presented in Figure 1E.3.7. At this point, the cells are ready for additional flow cytometric analysis or culture (see Basic Protocol, step 17)

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Knock-out culture medium 500 ml K-DMEM (Invitrogen, cat. no. 10829) 50 ml Knockout Serum Replacement (Invitrogen, cat. no. 10828; 10% v/v final) 10 ml 200 mM L-glutamine (Cellgro, cat. no. 25-005-CI) 5 ml antibiotic-antimycotic solution (Cellgro, cat. no. 30-004-CI) 10 ng/ml human recombinant epidermal growth factor (EGF; Sigma, cat. no. E9644) Store up to 15 days at 4◦ C Percoll solution, 24% As instructed by the manufacturer, prepare the 100% Percoll by mixing 9 parts pure Percoll (Sigma; cat. no. P-4937) with 1 part 10× HBSS (Invitrogen, cat. no. 14065056). Mix 12 ml of this 100% Percoll with unsupplemented DMEM medium in a final volume of 50 ml. Store the remaining 100% Percoll up to 4 weeks at 4◦ C.

Pre-digestion buffer Prepare 1000 ml calcium- and magnesium-free HBSS (CMF-HBSS; Lonza, cat. no. 04-315Q) supplemented with 0.5 mM EGTA (Sigma, cat. no. E4378). Store up to 4 weeks at 4◦ C.

Standard culture medium 430 ml DMEM (Invitrogen, high-glucose formulation; cat. no. 11960044) 5 ml 100 mM (100×) sodium pyruvate (Invitrogen, cat. no. 11360-070) 50 ml heat-inactivated fetal bovine serum (Invitrogen, cat. no. 16141; 10% v/v final) 5 ml 100 mM (100×) nonessential amino acids (Invitrogen, cat. no. 11140-050) 5 ml 200 mM L-glutamine (Cellgro, cat. no. 25-005-CI) 500 μl 55 mM 2-mercaptoethanol (Invitrogen, cat. no. 21985-023) 5 ml antibiotic-antimycotic solution (Cellgro, cat. no. 30-004-CI) 10 ng/ml human recombinant epidermal growth factor (EGF; Sigma, cat. no. E9644) Store up to 15 days at 4◦ C Transportation medium Prepare calcium- and magnesium-free HBSS (calcium- and magnesium-free; CMFHBSS; Lonza cat. no. 04-315Q) supplemented with antibiotic-antimycotic solution (Cellgro, cat. no.30-004-CI) according to the manufacturer’s instructions. Store up to 15 days at 4◦ C Isolation of Amniotic Epithelial Stem Cells

Final concentrations of antibiotics/antimycotics are 100 U/ml penicillin G, 100 μg/ml streptomycin, and 0.25 μg/ml amphotericin B.

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COMMENTARY Background Information Amniotic epithelial cells develop from the epiblast by 8 days after fertilization and prior to gastrulation. This raises the possibility that these cells might maintain the plasticity of pregastrulation embryo cells. The authors have determined that hAECs isolated from human term placenta express surface markers normally present on embryonic stem and germ cells (Miki et al., 2005). Isolated hAECs express SSEA-3 (8.79 ± 2.84%), SSEA-4 (43.94 ± 14.8%), TRA 1-60 (9.82 ± 4.31%), and TRA 1-81 (9.91 ± 4.49%), and do not express SSEA-1. In addition, hAECs express the pluripotent stem cell–specific transcription factors Oct-4, Sox-2, and Nanog. Under certain culture conditions, hAECs form spheroid structures that retain stem cell characteristics. hAECs do not require other cell-derived feeder layers to maintain Oct-4 and Nanog expression, do not express telomerase, display a normal karyotype, and are nontumorigenic upon transplantation (Miki and Strom, 2006). These characteristics indicate that hAECs are similar but not identical to embryonic stem cells. Based on their frequency in the tissue and their pluripotency, hAECs are also unlike adult tissue-specific stem cells.

Critical Parameters and Troubleshooting There is some variability in the cell yield between individuals. The authors ascribe these differences to the quality of tissue that is received, but this has not been specifically tested. It is clear that the time between the delivery of the placenta and the start of the cell isolation is critical. The tissue should be transferred to a refrigerator as soon as possible. This time in the refrigerator is called the cold ischemic time. It is best to minimize the cold ischemic time to <3 hr, if possible. Successful cell isolations have been conducted even 24 hr following delivery of the placenta, but the cell yield suffers considerably, and may be reduced to 1/3 of normal. The initial viability is not always indicative of the long-term quality of the cells. In many cases the initial viability is >75%, and yet the cells do not attach well to culture dishes and do not proliferate robustly in culture. The best estimate of long-term viability of the cells seems to be the plating efficiency, followed by the growth of the cells in the presence of EGF. If the initial plating efficiency of the

cells on tissue culture plastic is not >75%, the cells may not be optimal and should be discarded. Useful preparations attach at a high rate and also proliferate robustly in culture. Until one gets a better feel for the behavior of the cells in culture, it is helpful to use only those preparations that provide cells with high viability (trypan-blue negative; UNIT 1C.3) and high attachment over the first 24 hr of culture, followed by robust proliferation. It must also be understood that in addition to poor quality of the tissue, extensive trypsinization of the amnion membrane to release cells for periods longer than recommended here can also lead to poor viability and low plating efficiency.

Anticipated Results The yield of amniotic epithelial cells from one term placenta is 80 to 300 × 106 cells, which will continue to proliferate to passage 6.

Time Considerations The total time from stripping of the tissue to the final cell pellet is ∼2 to 2.5 hr.

Literature Cited Elwan, M.A. and Sakuragawa, N. 2007. Uptake and decarboxylation of L-3,4dihydroxyphenylalanine in cultured monkey placenta amniotic epithelial cells. Placenta 28:245-248. Elwan, M.A., Ishii, T., and Sakuragawa, N. 2003a. Characterization of dopamine D2 receptor gene expression and binding sites in human placenta amniotic epithelial cells. Placenta 24:658-663. Elwan, M.A., Ishii, T., and Sakuragawa, N. 2003b. Evidence of dopamine D1 receptor mRNA and binding sites in cultured human amniotic epithelial cells. Neurosci. Lett. 344:157-160. Kakishita, K., Elwan, M.A., Nakao, N., Itakura, T., and Sakuragawa, N. 2000. Human amniotic epithelial cells produce dopamine and survive after implantation into the striatum of a rat model of Parkinson’s disease: A potential source of donor for transplantation therapy. Exp. Neurol. 165:2734. Kakishita, K., Nakao, N., Sakuragawa, N., and Itakura, T. 2003. Implantation of human amniotic epithelial cells prevents the degeneration of nigral dopamine neurons in rats with 6-hydroxydopamine lesions. Brain Res. 980:4856. Miki, T. and Strom, S.C. 2006. Amnion-derived pluripotent/multipotent stem cells. Stem Cell Rev. 2:133-142. Miki, T., Lehmann, T., Cai, H., Stolz, D.B., and Strom, S.C. 2005. Stem cell characteristics of amniotic epithelial cells. Stem Cells 23:15491559.

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Sakuragawa, N., Thangavel, R., Mizuguchi, M., Hirasawa, M., and Kamo, I. 1996. Expression of markers for both neuronal and glial cells in human amniotic epithelial cells. Neurosci. Lett. 209:9-12. Sakuragawa, N., Misawa, H., Ohsugi, K., Kakishita, K., Ishii, T., Thangavel, R., Tohyama, J., Elwan, M., Yokoyama, Y., Okuda, O., Arai, H., Ogino, I., and Sato, K. 1997. Evidence for active acetylcholine metabolism in human amniotic epithelial cells: Applicable to intracerebral allografting for neurologic disease. Neurosci. Lett. 232:53-56. Takashima, S., Ise, H., Zhao, P., Akaike, T., and Nikaido, T. 2004. Human amniotic epithelial cells possess hepatocyte-like characteristics and functions. Cell Struct. Funct. 29:73-84. Wei, J.P., Zhang, T.S., Kawa, S., Aizawa, T., Ota, M., Akaike, T., Kato, K., Konishi, I., and Nikaido, T. 2003. Human amnion-isolated cells normalize blood glucose in streptozotocin- induced diabetic mice. Cell Transplant. 12:545552.

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Isolation and Manipulation of Mouse Trophoblast Stem Cells

UNIT 1E.4

Emi Himeno,1 Satoshi Tanaka,1 and Tilo Kunath2 1

Laboratory of Cellular Biochemistry, Animal Resource Sciences/Veterinary Medical Sciences, The University of Tokyo, Tokyo, Japan 2 MRC Centre for Regenerative Medicine, Institute for Stem Cell Research, University of Edinburgh, Edinburgh, United Kingdom

ABSTRACT The isolation of stable trophoblast stem (TS) cell lines from early mouse embryos has provided a useful cell culture model to study trophoblast development. TS cells are derived from pre-implantation blastocysts or from the extraembryonic ectoderm of early post-implantation embryos. The derivation and maintenance of mouse TS cells is dependent upon continuous fibroblast growth factor (FGF) signaling. Gene expression analysis, differentiation in culture, and chimera formation show that TS cells accurately model the mouse trophoblast lineage. This unit describes how to derive, maintain, and manipulate TS cells, including DNA transfection and chimera formation. Curr. Protoc. C 2008 by John Wiley & Sons, Inc. Stem Cell Biol. 7:1E.4.1-1E.4.27.  Keywords: trophoblast stem cells r TS cells r extraembryonic ectoderm r trophectoderm r trophoblast r FGF4

INTRODUCTION Trophoblast stem (TS) cells are stable, multipotent cell lines that can be derived from different stages of early mouse development (Tanaka et al., 1998). They are capable of producing all trophoblast cell types in culture and in vivo. In contrast to embryonic stem (ES) cells (UNIT 1C.4), TS cells contribute exclusively to the trophoblast lineage in chimeras, and do not colonize the embryo proper or the extraembryonic mesoderm lineages. TS cells are also incapable of producing extraembryonic endoderm cell types, such as visceral or parietal endoderm. These strict lineage restrictions, and other properties, such as maintenance of imprinted X-inactivation (Mak et al., 2002), have made TS cells a powerful and useful model for studying the mouse trophoblast lineage. This unit will describe derivation of TS cells from blastocysts and early post-implantation mouse embryos (Basic Protocol 1 and Alternate Protocol 1) and the maintenance of stem cells in culture (Basic Protocol 2). TS cells have been difficult to transfect by standard methods. Here, we describe a modified lipofection procedure (Basic Protocol 3), as well as two Alternate Protocols that work with TS cells. Finally, to test the developmental potential of TS cells, we briefly describe the production of TS cell chimeras (Basic Protocol 4).

STRATEGIC PLANNING The mouse strain from which the blastocysts are obtained for TS cell isolation is an important consideration. TS cell lines have been successfully derived from 129 strains, and outbred strains, such as ICR, CD-1, and PO (Pathology Oxford). However, derivation of TS cells from C57BL/6 has been problematic. Before embarking upon the task of deriving TS cell lines a stock of inactivated mouse embryonic fibroblasts (MEFs) should be generated, as well as MEF-conditioned medium (see Support Protocols 1 and 2). The

Current Protocols in Stem Cell Biology 1E.4.1-1E.4.27 Published online October 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e04s7 C 2008 John Wiley & Sons, Inc. Copyright 

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strain from which the MEFs are derived is not critical. MEFs from 129 or outbred mouse strains, such as CD-1 are suitable for derivation of TS cell lines. It should be carefully noted that it can take 3 to 4 months to derive stable TS cell lines suitable for downstream experiments. NOTE: All equipment and solutions coming into contact with live cells must be sterile and aseptic technique should be used accordingly. NOTE: All incubations at 37o C are performed in a humidified 5% CO2 /95% air incubator unless otherwise stated. NOTE: All centrifugations are performed at room temperature unless otherwise stated. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations for the care and use of laboratory animals. BASIC PROTOCOL 1

ISOLATION OF TROPHOBLAST STEM CELLS FROM BLASTOCYSTS The derivation of TS cell lines from mouse blastocysts is similar to ES cell derivation. The procedures for flushing embryos, plating on mouse embryonic fibroblasts (MEFs), and disaggregating blastocyst outgrowths are all similar. However, the critical difference is that mouse TS cells require exogenous FGF4 in the culture medium while mouse ES cells require LIF.

Materials Mitomycin C–treated MEFs (MMC-MEFs; see Support Protocol 1) TS medium (see recipe) TS + F4H medium (see recipe) 3.5 days post-coitum (dpc) mice M2 medium (Sigma, cat. no. M7167) KSOM medium (optional) Acid Tyrode’s (optional) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.1% (w/v) trypsin/1 mM EDTA (see recipe) 70CM + 1.5× F4H medium (see recipe and Support Protocol 2) TS + 1.5× F4H medium (see recipe) 4-well and 6-well tissue culture plates 100-mm petri dishes 1-ml syringes and 26-G needles Dissecting microscope Finely-drawn mouth (Pasteur) pipets with tubing and mouthpiece Inverted microscope Organ culture dish (optional) 20-μl and 200-μl adjustable pipets U-bottom 96-well nontissue culture plates 35-mm tissue culture dishes (optional) Additional reagents and equipment for euthanasia of mice (Donovan and Brown, 2006) Isolation and Manipulation of Mouse Trophoblast Stem Cells

Prepare 4-well plates of MEFs 1. Day −1: The day before blastocyst collection, plate MMC-MEFs in 4-well plates at 4 × 104 cells in 0.5 ml TS cell medium per well. The plates do not need to be coated with gelatin.

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Although 24-well plates have a similar surface area per well as 4-well plates (2 cm2 versus 1.4 cm2 ), their use is not advised because the high walls of 24-well plates make disaggregation of blastocyst outgrowths difficult.

2. Day 0: On the day of blastocyst collection, add FGF4 and heparin to a sufficient volume of TS cell medium to make TS + F4H medium (i.e., 2 ml per 4-well plate). Replace the medium in all 4-well plates from TS cell medium with 0.5 ml TS + F4H medium per well. If necessary due to time constraints, MMC-MEFs may be plated a few hours before blastocyst collection directly in TS + F4H medium, instead of the day before.

Collect blastocysts 3. Day 0: Sacrifice 3.5 dpc pregnant mice by cervical dislocation (Donovan and Brown, 2006). Dissect out uterine horns and place in a drop of M2 medium in an inverted lid from a 100-mm petri dish. Fill a 1-ml syringe with M2 medium, attach a 26-G needle, and flush blastocysts from the uterine horns (Nagy et al., 2003). This procedure is normally performed on the bench. It can be performed in a laminar flow hood if additional sterility is desired.

4. Under a dissecting microscope, use a finely-drawn mouth pipet to collect and pool all the blastocysts into a fresh drop of M2 medium. Transfer the embryos through several drops of M2 medium until all the maternal tissue and blood has been removed. 5. Using a finely-drawn mouth pipet and an inverted microscope place one blastocyst in each well of the 4-well plates (containing MMC-MEFs and TS + F4H medium) and incubate at 37◦ C. A flame (e.g., alcohol burner) can be placed near the microscope to increase sterility, or the procedure can be performed in a laminar flow hood if the microscope can be placed within it. If more than expected blastocyst numbers are obtained, plate the most advanced looking embryos in the prepared 4-well plates, and transfer the additional embryos into an organ culture dish with KSOM medium. Plate MMC-MEFs in additional 4-well plates in TS + F4H medium (0.5 ml per well), and then add the remaining blastocysts anytime after the MEFs have attached (from 1 hr later to the next day). Do not plate morula or nascent blastocysts—allow these embryos to mature in KSOM overnight before transferring to 4-well plates.

6. Day 1: Monitor blastocysts for hatching from the zona pellucida and attachment to the MEFs. Do not feed the cultures. If the blastocysts have not hatched by Day 2, they may be hatched with acid Tyrode’s (Nagy et al., 2003).

7. Day 2: If the embryos are firmly attached and have formed an outgrowth on the second day, carefully aspirate the medium and feed with 0.5 ml freshly prepared TS + F4H medium per well. If the blastocysts have not attached yet, wait an additional 2 to 3 days for attaching to occur. If the outgrowth is very loosely attached, remove half of the medium (250 μl) and carefully feed with TS + F4H (250 μl).

Disaggregate blastocyst outgrowths 8. Days 3 to 5: Perform disaggregation of the blastocyst outgrowths on days 3, 4, or 5 when the outgrowth is ∼800 to 1000-μm in diameter. Carefully aspirate the medium and wash the outgrowth twice, each time with 0.5 ml CMF-PBS. The efficiency of TS cell line derivation decreases if the outgrowth is too big.

9. Add 0.1 ml 0.1% trypsin/1 mM EDTA, then incubate 5 min at 37◦ C.

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10. Thoroughly disaggregate the outgrowth by pipetting with a 200-μl adjustable pipet set at ∼95 μl. This is performed in a tissue culture hood without the aid of a microscope. Crudely pipetting up and down the entire contents of the well is usually sufficient to achieve disaggregation. A finely-drawn mouth pipet under a microscope may be used for this step, but an adjustable pipet is recommended if disaggregating many outgrowths. Change tips between outgrowths to avoid cross-contamination of cells.

11. Add 0.5 ml 70CM + 1.5× F4H medium to stop trypsinization. Culture 8 to 16 hr at 37◦ C, then change the medium and add 0.5 ml fresh 70CM + 1.5× F4H medium. Hereafter, change the 70CM + 1.5× F4H medium every other day. Tight epithelial TS cell colonies will become apparent 3 to 7 days after step 11 is completed (Fig. 1E.4.1A,B). ES cell colonies very rarely appear in these conditions. The day of appearance can be very variable and can depend on the mouse strain used. The appearance of parietal endoderm cells is fairly common. They will not form epithelial colonies, and will appear as highly refractile, bright individual cells (Fig. 1E.4.1C).

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Figure 1E.4.1 Trophoblast stem cell cultures. (A) A small TS cell colony on MEFs. (B) A highmagnification picture of the edge of a TS cell colony (upper-left) to illustrate the raised edge of the colony. MEFs and some parietal endoderm cells are present. (C) Parietal endoderm cells that may contaminate TS cell cultures. (D) A TS cell colony grown on tissue culture plastic with 70CM + F4H in the absence of MEFs. (E) Differentiated giant cells, 4 days after the removal of FGF4 from the cell culture medium. (F) A higher magnification of trophoblast giant cells derived from TS cells. The inset shows a single giant cell (left) beside an undifferentiated TS cell colony (right).

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Isolate TS cell colonies If extraembryonic endoderm cells are not present, the following isolation steps (12 to 19) may be omitted. 12. Days 6 to 12: Prepare 4-well plates by plating 4 × 104 cells of MMC-MEFs per well in 0.5 ml TS + 1.5× F4H medium at least 1 hr before proceeding to step 13. 13. If possible, place the dissecting microscope within a tissue culture hood. In a Ubottom nontissue culture 96-well plate, place 50-μl aliquots of 0.1% trypsin/1 mM EDTA in the number of wells equivalent to the number of TS cell colonies to be isolated. Encircling the TS colonies using an inverted microscope on the bottom surface of 4-well plates with a fine marker will aid with the picking of TS colonies under the dissecting microscope.

14. Aspirate the medium from the 4-well plates containing TS cell colonies and unwanted cells, wash the cells once with 0.5 ml CMF-PBS, then add 0.3 ml CMF-PBS. Do not remove this last CMF-PBS wash. 15. Under the dissecting microscope, pick TS cell colonies by using a 20-μl adjustable pipet (or a finely-drawn glass pipet) containing a small amount (∼5 μl) of 0.1% trypsin/1 mM EDTA. Put the picked TS colonies into the U-bottom wells containing trypsin/EDTA solution (prepared in step 13). 16. Once all colonies are picked up, incubate the 96-well plate 5 min at 37◦ C. 17. Add 150 μl TS + 1.5× F4H medium to stop trypsinization and separate the cells by thorough pipetting. Transfer the entire contents to the 4-well plates containing fresh MMC-MEFs in 0.5 ml TS + 1.5× F4H medium (prepared in step 12). 18. Incubate 8 to 16 hr at 37◦ C. After the incubation, exchange the medium with 0.5 ml fresh TS + 1.5× F4H and continue to culture. 19. After 2 days, change the medium every other day with 0.5 ml TS + F4H medium per well. TS cell colonies should re-emerge 3 to 7 days after isolation.

Passage TS cells 20. When the colonies appear over-grown or the well is about half-confluent, passage TS cells with trypsin to a 35-mm dish or 6-well plate of preplated MMC-MEFs (see Table 1E.4.1 for MEF numbers) in 2 ml TS + F4H medium. 21. Two days after step 20 is completed, replace the medium with 2 ml fresh TS + F4H medium. Table 1E.4.1 Recommended Number of MMC-MEFs for Co-Culturing with TS Cells and Suggested Volumes of TS Medium and Trypsin Solution for Routine Culture

Number of MMC-MEFs for coculture

Medium (ml)

0.05% trypsin/ 1 mM EDTA (ml)

150-mm

3 × 106

25

5

100-mm

1.2 × 10

10

2

60-mm

4 × 10

5

1

35-mm

2 × 10

2

0.5

4 well

4 × 10 /well

0.5

0.1

Plate

6

5 5

4

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22. Keep changing the medium (2 ml of TS + F4H) every other day. Gradually expand culture of TS cells to larger dishes or flasks. Note that significant differentiation occurs for the first several passages. TS cells may be frozen for storage once they are in 60-mm dishes.

ALTERNATE PROTOCOL 1

ISOLATION OF TROPHOBLAST STEM CELLS FROM POST-IMPLANTATION EMBRYOS The derivation of TS cell lines from extraembryonic ectoderm (ExE) or chorionic ectoderm (ChE) of post-implantation embryos may greatly reduce the chance of extraembryonic endoderm contamination. Handling skills for working with post-implantation embryos, however, will be required. The protocol described here is for 6.5 dpc embryos. The derivation of TS cell lines from 7.5 dpc ChE is similar and is described in a paper by Uy and colleagues (Uy et al., 2002).

Materials Mitomycin C-treated MEFs (MMC-MEFs; see Support Protocol 1) 0.5% (w/v) trypsin/2.5% (w/v) pancreatin (see recipe) 6.5 dpc mice Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) CMF-PBS/10% (v/v) FBS 0.1% (w/v) trypsin/1 mM EDTA (see recipe) TS + F4H medium (see recipe) 60-mm Center-well Organ Culture Dish (BD Falcon, cat. no. 353037) 100-mm petri dishes 200-μl pipet tips Fine forceps Dissecting microscope Glass needles or 29-G needles Finely-drawn mouth (Pasteur) pipets with tubing and mouthpiece U-bottom 96-well nontissue culture plates Inverted microscope Prepare 4-well plates of MEFs 1. Day −1: Prepare 4-well plates of MMC-MEFs (Basic Protocol 1, steps 1 and 2). Collect extraembryonic ectoderm from 6.5 dpc embryos 2. Day 0: Thaw a tube of 0.5% trypsin/2.5% pancreatin (1-ml aliquot) on ice. Briefly centrifuge at 10,000 × g, 4◦ C to remove debris, and transfer the supernatant into the center well of an organ culture dish and keep on ice. 3. Dissect out 6.5 dpc embryos in CMF-PBS in 100-mm petri dishes, then transfer the embryos into CMF-PBS/10% FBS using a cut-off 200-μl pipet tip (Nagy et al., 2003). 4. Remove Reichert’s membrane with a pair of fine forceps under a dissecting microscope.

Isolation and Manipulation of Mouse Trophoblast Stem Cells

5. Cut off the ectoplacental cone and the embryonic part of the embryos (Fig. 1E.4.2A,B) by using a pair of glass needles or 29-G needles. Transfer “extraembryonic pieces” (Ex in Fig. 1E.4.2B) to CMF-PBS/10% FBS in the center well of an organ culture dish.

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Figure 1E.4.2 Collection of extraembryonic ectoderm from 6.5 dpc embryo. Before (A) and after (B) separation of a 6.5 dpc embryo into an ectoplacental cone (EPC), an extraembryonic piece (Ex) consisting of visceral endoderm (VE) and extraembryonic ectoderm (ExE), and an embryonic piece (Em) consisting of VE and epiblast. Ex was further separated into VE and ExE by trypsin/pancreatin treatment followed by pipetting (C). Lines in A indicate two excisions.

Separate VE and ExE 6. Transfer the extraembryonic pieces into 0.5% trypsin/2.5% pancreatin and incubate 20 min at 4◦ C or on ice. 7. Transfer the extraembryonic pieces back to the organ culture dish containing CMFPBS/10% FBS. 8. Separate the layer of visceral endoderm (VE) from that of extraembryonic ectoderm (ExE) by pipetting through a finely-drawn Pasteur pipet. The width of this drawn pipet should be only slightly larger than the tissue being manipulated. The ExE has a relatively clear appearance, whereas the VE is darker under a dissecting microscope (Fig. 1E.4.2C).

9. Collect ExE pieces in CMF-PBS/10% FBS in a new organ culture dish. Discard the VE pieces. If derivation of TS cell lines from genetically distinct embryos is required, put each ExE piece into wells of 4-well plates and use the corresponding pieces of VE for genotyping.

Dissociate ExE pieces 10. With a cut-off 200-μl pipet tip, transfer ExE pieces into CMF-PBS in an organ culture dish to rinse away the FBS, and then transfer them individually into an empty well of a U-bottom 96-well nontissue culture plate along with 40 to 50 μl CMF-PBS. ExE pieces become very sticky in CMF-PBS.

11. Add 40 μl 0.1% trypsin/1 mM EDTA to each well containing ExE pieces, and gently pipet to mix. Incubate 8 min at 37◦ C. Uy et al. used pronase and EGTA instead of trypsin/EDTA, as has been described for dissociation of epiblast (Gardner and Davies, 2000; Uy et al., 2002).

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12. Stop trypsinization by adding 80 μl TS + F4H medium. Disaggregate the cells by pipetting ∼30 times through a 200-μl tip.

Plate and grow cells 13. Using an inverted microscope and a P200 pipet with a 200-μl tip, transfer entire contents to one well of the 4-well plate containing preplated MMC-MEFs and 0.5 ml TS + F4H medium (step 1). Add one ExE per well. 14. Day 1: Remove medium and re-feed with 0.5 ml freshly prepared TS + F4H medium per well. Hereafter, change the medium every other day. 15. Follow steps 20 to 22 of Basic Protocol 1 to passage TS colonies. Because only a small amount of non-TS cells are expected to appear in this procedure, elimination of such contaminated non-TS cells by mechanically lifting them is not likely to be necessary. SUPPORT PROTOCOL 1

MITOMYCIN C TREATMENT OF MOUSE EMBRYONIC FIBROBLASTS TS cells require soluble factors secreted from MEFs to maintain their undifferentiated, proliferative status. It is now known that TGFβ and activin are the critical components provided by the MEFs (Erlebacher et al., 2004). However, we continue to use MEFs because they are relatively cheap and they may be providing additional noncritical factors beneficial to TS cells. To coculture TS cells with MEFs, mitotic inactivation is needed to prevent their overgrowth. Mitomycin C treatment of MEFs, which irreversibly inhibits DNA synthesis by producing interstrand DNA cross-links, is used for this purpose. Irradiated MEFs have also been successfully used for TS cell derivation and maintenance.

Materials Mouse embryonic fibroblasts (MEFs), frozen DMEM/10% FBS (see recipe) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.05 % (w/v) trypsin/1 mM EDTA (see recipe) Mitomycin C (MMC; see recipe) 2× freezing medium for MEFs (see recipe) 37◦ C water bath 50-ml centrifuge tubes 150-mm tissue culture dishes Freezing vials Cell-freezing container (e.g., 5100 Cryo 1◦ C Freezing Container, Nalgene) Liquid nitrogen tank Additional reagents and equipment for performing a viable cell count (UNIT 1C.3) Thaw MEFs 1. Thaw a frozen vial of MEFs in a 37◦ C water bath. A vial contains primary MEFs harvested from an 80% to 90 % confluent 150-mm dish (∼2.5 × 106 cells). Ideally, MEFs should be as low passage as possible.

2. Transfer entire contents into a 50-ml tube containing 10 ml DMEM/10% FBS and centrifuge 3 min at 200 × g. 3. Discard supernatant and resuspend the cells gently in 25 ml DMEM/10% FBS. Isolation and Manipulation of Mouse Trophoblast Stem Cells

Plate cells 4. Split cells (5 ml each) into five 150-mm dishes, each containing 20 ml DMEM/10% FBS.

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Table 1E.4.2 Recommended Number of MMC-MEFs for Preparation of Conditioned Medium to be Used in MEF-Free Culture of TS Cells

Plate

Number of MMC-MEFs

TS medium (ml)

150-mm

6 × 106

25-27

100-mm

2.4 × 10

6

10-12

5. Culture cells at 37◦ C to ∼90% confluency (3 to 4 days).

Passage cells 6. Remove the medium and rinse twice, each time with 10 ml CMF-PBS per dish. 7. Add 5 ml 0.05% trypsin/1 mM EDTA to each dish and incubate 3 min at 37◦ C. 8. Add 10 ml DMEM/10% FBS to each dish and pipet gently to break cell aggregates. 9. Transfer cell suspensions to 50-ml tubes and centrifuge 3 min at 200 × g. 10. Discard supernatant and resuspend the cells in a 50-ml tube containing 40 ml DMEM/10% FBS. 11. Split cells (2 ml each) into twenty 150-mm dishes, each containing 23 ml DMEM/10% FBS.

Treat cells with mitomycin C 12. Culture cells at 37◦ C to ∼90% confluency (2 to 3 days). Change the medium on the second day (optional). 13. When the cells become almost confluent, remove the medium and add 10 ml DMEM/10% FBS containing 10 μg/ml MMC. One vial of MMC from Sigma-Aldrich contains 2 mg, making it convenient to prepare an exact volume of MMC-containing medium for twenty 150-mm dishes. If desired, this production may be scaled down for five or ten 150-mm dishes.

14. Incubate cells 2 hr at 37◦ C . 15. Remove the medium and rinse cells twice, each time with 10 ml CMF-PBS. 16. Trypsinize cells as described in steps 6 to 9. Finally, resuspend the cell pellet in 40 ml DMEM/10% FBS. 17. Count viable cells (UNIT 1C.3) and dilute to 2× of final desired cell density with DMEM/10% FBS. The final desired cell density may vary depending on the frequency and the scale of TS cell culture. For example, we make aliquots at three different densities, i.e., 6, 2.4 and 1.2 × 106 cells/ml/vial, which we thaw for TS cell coculture in five, two, and one 100-mm plate(s), respectively (see Tables 1E.4.1 and 1E.4.2).

Freeze MEFs 18. Add an equal volume of 2× freezing medium for MEFs and mix gently. 19. Add 1 ml of cell suspension to each freezing vial, and store at –80◦ C in a cell-freezing container overnight. 20. The next day, transfer the vials to a liquid nitrogen tank. Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 2

PREPARATION OF MEF-CONDITIONED MEDIUM Maintenance of TS cells is dependent on exogenous FGF4 and soluble factors secreted from MEFs. During the course of TS cell derivation from blastocysts, MMC-MEFs will be continuously cultured for >1 week, going through trypsinization and replating steps, which may damage the inactivated-MEFs and may hamper their ability to produce critical factors. MEF-conditioned medium (MEF-CM) is used to compensate for such potential under-performance of MEFs. MEF-CM is also used when TS cells are maintained on tissue culture plastic in MEF-free conditions, for example, in 70CM + F4H medium.

Materials Mitomycin C–treated MEFs (MMC-MEFs; see Support Protocol 1) DMEM/10% FBS (see recipe) TS medium (see recipe) 37◦ C water bath 50-ml centrifuge tubes 100-mm dishes or 150-mm dishes 0.22-μm filter unit for a glass bottle (Millipore) Glass fiber prefilter (Millipore) 500-ml glass bottles, autoclaved 1. Thaw a frozen vial of MMC-MEFs cells quickly in a 37◦ C water bath. See Table 1E.4.2 for the appropriate cell numbers to each culture dish.

2. Add the cells to 10 ml DMEM/10% FBS in a 50-ml tube and centrifuge 3 min at 200 × g. 3. Discard the supernatant. 4. Resuspend the cells in TS medium (without FGF4 and heparin) and seed in 150-mm or 100-mm tissue culture dishes. Use 25 to 27 ml TS medium per 150-mm dish or 10 to 12 ml/10-mm dish. Use TS medium without penicillin and streptomycin if preparing MEF-CM to be used during lipofection (Basic Protocol 3).

5. Incubate cells 3 days at 37◦ C without changing the medium. 6. Collect the medium in 50-ml tubes and store at −20◦ C while preparing additional batches. Prepare two more batches with the same dish of MMC-MEFs. In total, three batches of MEF-CM are collected over 9 days.

7. Thaw and pool all three batches of MEF-CM. Centrifuge 20 min at 2300 × g, 4◦ C, to remove debris. 8. Collect the supernatant and filter through a 0.22-μm filter with a glass fiber prefilter into 500-ml glass bottles. 9. Store at −20◦ C in 30- to 40-ml aliquots. Thaw each aliquot as needed and store up to 1 month at 4◦ C; do not refreeze. Alternatively, MEF-CM may be immediately spun and filtered upon collection, then aliquoted and frozen. Isolation and Manipulation of Mouse Trophoblast Stem Cells

1E.4.10 Supplement 7

Current Protocols in Stem Cell Biology

MAINTENANCE OF TS CELLS TS cells are virtually immortal and have been expanded for >50 passages under appropriate conditions with no apparent change in their morphology or viability. Established TS cells can be passaged at 1:10 to 1:20 every 4 to 6 days. The karyotype of TS cells is predominantely diploid. However, tetraploid cells are often present, consistent with differentiated cells, and some translocations have been identified (Uy et al., 2002). This has not affected the ability of TS cells to differentiate or contribute to chimeras.

BASIC PROTOCOL 2

Materials TS cells in culture Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.05% (w/v) trypsin/1 mM EDTA (see recipe) TS medium (see recipe) TS medium + F4H (see recipe) 70CM + F4H medium (optional; see recipe) 50-ml centrifuge tubes Cell culture dishes (see Table 1E.4.1 for sizes) 1. When the TS cells reach ∼80% confluency, aspirate the medium and rinse twice, each time with CMF-PBS (e.g., 10 ml for 100-mm dish). 2. Add 0.05% trypsin/1 mM EDTA to the dish and incubate 3 min at 37◦ C. See Table 1E.4.1 for appropriate volume.

3. Add TS medium to stop the reaction and disaggregate cell aggregates by gentle pipetting. Note that differentiated cells are more resistant to trypsin than true TS cells. Therefore, steps 2 and 3 should not be performed too aggressively.

4. Transfer the cell suspension to a 50-ml tube and centrifuge 3 min at 200 × g. 5. Discard supernatant and resuspend the cells with an appropriate volume of TS medium. 6. Transfer 1/10 to 1/20 of the cell suspension to a new dish of MMC-MEFs in TS + F4H medium (see Table 1E.4.1 for appropriate cell number and size of dish) and culture at 37◦ C. Feed TS cells with TS + F4H medium every other day and passage when cells reach ∼80% confluency (4 to 6 days). Use 70CM + F4H medium under MEF-free conditions.

REMOVING MMC-MEFs FROM TS CELL CULTURES Removal of MMC-MEFs from TS culture may be required for DNA/RNA/protein extraction from TS cells, induction of differentiation, or DNA transfection (see Basic Protocol 3). MEFs and differentiated trophoblast cells adhere to the tissue culture dish more quickly than TS cells. This differential plating time can be used to recover floating TS cells in the medium after the MEFs and other cell types have adhered to the tissue culture plastic. TS cells can be maintained in the absence of MMC-MEFs in medium supplemented with 70% MEF-conditioned medium (see Support Protocol 2). The example below is for a 100-mm cell culture dish. Adjust volumes accordingly for different sizes of dishes or flasks.

Materials Cultures of TS cells Mitomycin C–treated MEFs (MMC-MEFs; see Support Protocol 1) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen)

SUPPORT PROTOCOL 3

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0.05% (w/v) trypsin/1 mM EDTA (see recipe) TS medium (see recipe) 70CM + F4H medium (see recipe) TS + F4H medium (see recipe) 100-mm cell culture dishes 50-ml centrifuge tubes Additional reagents and equipment for performing a viable cell count (UNIT 1C.3) 1. Grow TS cells on MMC-MEFs to ∼80% to 90% confluency in a 100-mm dish. 2. Discard the medium and rinse twice, each time with 10 ml CMF-PBS. 3. Add 1 ml 0.05% trypsin/1 mM EDTA and incubate 3 min at 37◦ C. 4. Add 9 ml TS medium and break up cell aggregates by gentle pipetting. 5. Transfer cell suspension to a 50-ml tube and centrifuge 3 min at 200 × g. 6. Discard supernatant and resuspend the cells in 10 ml 70CM + F4H medium. 7. Transfer the suspension to a 100-mm culture dish and incubate 45 to 60 min at 37◦ C. 8. Carefully collect the medium containing floating TS cells and plate in a new 100-mm culture dish. Count viable cells (UNIT 1C.3) before plating if needed. Approximately 5 × 105 TS cells per 100-mm culture dish will reach 80% to 90% confluency in 3 to 4 days under MEF-free conditions. The first 100-mm culture dish (from step 7) may be discarded or TS + F4H medium (10 ml) may be added and cells cultured at 37◦ C to recover additional TS cell colonies. SUPPORT PROTOCOL 4

FREEZING TS CELLS TS cells can be frozen at a lower density than ES cells. For example, TS cells from an ∼80% confluent 100-mm dish can be divided into nine cryovials, each of which is sufficient to be replated in a single 100-mm dish.

Materials 2× freezing medium for TS cells (see recipe) TS cell cultures TS medium (see recipe) 1-ml cryovials Cell-freezing container (e.g., 5100 Cryo 1◦ C Freezing Container, Nalgene) −80◦ C freezer Liquid nitrogen tank Additional reagents and equipment for trypsinization and pelleting of cells (Basic Protocol 2) 1. Prepare 2× freezing medium for TS cells and keep on ice. 2. Harvest TS cells from an ∼80% confluent culture by trypsinization and pellet cells by centrifugation (see Basic Protocol 2, steps 1 to 4). 3. Discard supernatant and resuspend the cells in TS medium (e.g., 4.5 ml for 100-mm dish) and add an equal volume of 2× freezing medium for TS cells. Isolation and Manipulation of Mouse Trophoblast Stem Cells

1E.4.12 Supplement 7

4. Mix gently and aliquot 1 ml of cell suspension per cryovial. 5. Put the cryovials into a cell-freezing container and place in a −80◦ C freezer overnight. 6. Transfer the cryovials to a liquid nitrogen tank the following day. Current Protocols in Stem Cell Biology

THAWING TS CELLS Frozen stocks of TS cells should be thawed using the following protocol. Thawing onto MMC-MEFs is better for cell viability and reduced differentiation.

SUPPORT PROTOCOL 5

Materials Mitomycin C–treated MEFs (MMC-MEFs; see Support Protocol 1) Vials of frozen TS cells TS medium (see recipe) TS + F4H medium (see recipe) 37◦ C water bath 50-ml centrifuge tubes 100-mm cell culture dishes 1. Prepare MMC-MEFs culture dish by plating them at the required density for coculture at least 1 hr before thawing TS cells (see Table 1E.4.1). 2. Thaw a frozen vial of TS cells quickly in a 37◦ C water bath. 3. Add thawed cells to 10 ml TS medium in a 50-ml tube and centrifuge 3 min at 200 × g. 4. Discard supernatant and tap the bottom of the tube gently to loosen the cell pellet. 5. Add an appropriate volume of TS + F4H medium (see Table 1E.4.1) and seed onto MMC-MEF plates prepared in step 1. 6. Change the medium the next day to remove cell debris. 7. Replace with fresh TS + F4H medium every 2 days. 8. Passage the cells as required (see Basic Protocol 2, steps 1 to 6).

GENETIC MANIPULATION OF TS CELLS This section describes three methods to genetically manipulate TS cells. All methods involve the introduction of exogenous DNA. Transfection with Lipofectamine is the most efficient (Basic Protocol 3), followed by Nucleofection (Alternate Protocol 2). If a single copy of the exogenous transgene is required, then electroporation is the best choice (Alternate Protocol 3). The establishment of stably transformed TS cell lines from any of the methods is also described (Support Protocol 6).

DNA Transfection with Lipofectamine Lipofectamine is one of the most useful and common transfection regents, but the efficiency of transfection into TS cells by the manufacturer’s protocol is very low (∼1%). Here, we introduce a more efficient method by using petri dishes, which keep TS cells floating during the transfection procedure. The efficiency of transfection improves to 20% to 30% using this protocol (Fig. 1E.4.3B through D).

BASIC PROTOCOL 3

Materials TS cells Mitomycin C–treated MEFs (MMC-MEFs; Support Protocol 1) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.05% (w/v) trypsin/1 mM EDTA (see recipe) TS medium (see recipe) TS medium + 1.5× F4H (see recipe) 70CM + 1.5× F4H medium (see recipe)

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Figure 1E.4.3 Transfection of TS cells. (A) TS cells on MEFs 3 days after passage. Small and uniform colonies should be prepared for effective transfection. (B) The expression of DsRed 24 hr after transfection. (C) TS colony after 14 days of drug selection. (D) DsRed expression in TS cells after a few passages. Scale bar 200 μm in A through D. (E, F) TS cell colonies after 12 days of neomycin selection. The colonies were fixed and stained with X-gal for β-galactosidase activity. One colony exhibited homogenous expression (E), while the other was more heterogeneous (F).

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Current Protocols in Stem Cell Biology

Plasmid DNA (4 μg) in 10 to 20 μl sterile H2 O (linearize plasmid for stable lines) Opti-MEM (Invitrogen) Lipofectamine 2000 (Invitrogen) 1 mM EDTA/CMF-PBS (see recipe) 50-ml tubes 1.5-ml microcentrifuge tubes 35-mm petri dishes or 6-well non-tissue culture plates 100-mm culture dishes Additional reagents and equipment for counting viable cells (UNIT 1C.3) Prepare TS cells 1. Grow TS cells on MMC-MEFs to ∼80% confluency in a 100-mm dish (Fig. 1E.4.3A). Approximately 3 to 5 × 105 TS cells are needed per transfection after removal of MMCMEFs. An ∼80% confluent 100-mm TS cell culture should yield 5 to 7 × 106 TS cells after removal of MMC-MEFs. Overgrowing TS cells cause a decreased efficiency of the transfection. Uniform TS cell colonies lead to high transfection efficiencies. A few passages may be needed to obtain TS cells in ideal conditions.

2. Discard the medium and rinse twice, each time with 10-ml CMF-PBS. 3. Add 2 ml 0.05% trypsin/1 mM EDTA and incubate 3 min at 37◦ C. 4. Add 8 ml TS medium and break cell aggregates by gently pipetting. 5. Transfer cell suspensions to 50-ml tubes and centrifuge for 3 min at 200 × g. 6. Discard supernatant and resuspend the cells in 10 ml TS + 1.5× F4H medium. 7. Transfer the suspension to a new dish and incubate 45 to 60 min at 37◦ C. This step removes MMC-MEFs. During this time, prepare Lipofectamine complex following steps 10 to 13.

8. Collect the supernatant slowly and count viable cells (UNIT 1C.3). 9. Prepare 5 × 105 cells/ml with 70CM +1.5× F4H medium in 50-ml tubes. If possible, use medium that does not contain penicillin/streptomycin to increase the viability of cells after transfection.

Prepare Lipofectamine complex 10. Dilute 4 μg plasmid DNA in 250 μl Opti-MEM in a 1.5-ml microcentrifuge tube. 11. In a separate 1.5-ml microcentrifuge tube, add 10 μl Lipofectamine 2000 to 250 μl Opti-MEM, mix gently by pipetting, and incubate for 5 min at room temperature. 12. Add the DNA mixture to the Lipofectamine 2000 mixture. 13. Mix gently by pipetting and incubate for 20 to 40 min at room temperature.

Transfect floating TS cells 14. Drop the Lipofectamine complex (∼510 to 520 μl) into an empty 35-mm petri dish or 6-well nontissue culture dish. 15. Add 1 ml cell suspension (5 × 105 cells/ml) to the Lipofectamine complex and mix well by gently pipetting. 16. Incubate 4 to 5 hr at 37◦ C.

Passage to culture dish 17. Transfer supernatant to a 50-ml tube.

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18. Add 1 ml of 1 mM EDTA/CMF-PBS to the dish and incubate 3 min at 37◦ C to strip cells adhered to the dish. 19. Add 1 ml TS medium to wash and collect cells into the 50-ml tube from step 17. 20. Centrifuge 3 min at 200 × g. 21. Discard supernatant and resuspend the cells in 10 ml of 70CM + 1.5× F4H medium (penicillin/streptomycin-free, if possible). 22. Seed all the cells from one transfected well to a 100-mm tissue culture dish and incubate 24 hr at 37◦ C. If a fluorescent marker is used, observe successfully transfected cells at this time (Fig. 1E.4.3B). 23. Discard medium and add medium with the appropriate antibiotic to select for the introduced plasmid. After 24 hr from transfection, use normal 70CM +1.5× F4H medium with penicillin and streptomycin, if desired. The following concentration of antibiotics work with TS cells: neomycin 100 to 200 μg/ml and zeocin 200 μg/ml.

24. Change the medium every second day. Passage TS cells in bulk or use them to establish clonal, stable cell lines (see Support Protocol 6). ALTERNATE PROTOCOL 2

Nucleofection of TS Cells This method of transfection is less efficient, but is also less labor-intensive than the Lipofectamine protocol. However, a nucleofector device is required. The protocol described below uses reagents originally designed for ES cells (Lakshmipathy et al., 2007).

Materials Mouse ES Cell Nucleofector Kit (Amaxa, cat no. VPH-1001) containing: Supplement Mouse ES Cell Nucleofector Solution TS cells 70CM + F4H medium (see recipe) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.05% (w/v) trypsin/1 mM EDTA (see recipe) TS medium (see recipe) Plasmid DNA (5 μg) in 1 to 5 μl sterile H2 O 50-ml tubes 15-ml tubes (optional) Amaxa-certified cuvette (included in the Mouse ES Nucleofector kit) Nucleofector II Device (Amaxa, cat no. AAD-1001) 100-mm culture dishes Additional reagents and equipment for counting viable cells (UNIT 1C.3) Prepare TS cells 1. Add 0.5 ml Supplement to 2.25 ml Mouse ES Cell Nucleofector Solution. This mixture is stable up to 3 months at 4◦ C.

2. Culture TS cells without MEFs in 70CM + F4H to ∼80% to 90% confluency. 3. Aspirate medium and rinse twice, each time with 10 ml CMF-PBS per 100-mm dish. Isolation and Manipulation of Mouse Trophoblast Stem Cells

4. Add 1 ml of 0.05% trypsin/1 mM EDTA per dish and incubate 3 min at 37◦ C. 5. Add 9 ml TS medium per dish and break cell aggregates to a single-cell suspension by gently pipetting.

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Current Protocols in Stem Cell Biology

6. Transfer cell suspension into 50-ml tubes and count a 25-μl of cells with a hemacytometer (UNIT 1C.3). 7. Transfer 1 to 2 × 106 cells into a new 50-ml tube (or 15-ml tube) and centrifuge 3 min at 200 × g. 8. Aspirate supernatant and resuspend the cell pellet in 100 μl of ES Cell Nucleofector Solution plus Supplement (prepared in step 1).

Nucleofect TS cells 9. Add DNA (5 μg) to TS cells in 100 μl of supplemented Nucleofector solution. There is some evidence that circular plasmid DNA may promote single integration sites more often than linear DNA for this protocol. However, the transfection efficiency is reduced with circular DNA.

10. Mix by pipetting up and down, then transfer to an Amaxa-certified cuvette included in the Mouse ES Nucleofector kit. 11. Insert cuvette into Nucleofector II Device, select program A-30, and press the X button to start the program. 12. Transfer the entire mixture to 10 ml prewarmed (37◦ C) 70CM + F4H and plate in a 100-mm culture dish. 13. Start drug selection on the second or third day after nucleofection. 14. Change the medium (70CM + F4H + drug) every second or third day until individual colonies appear (7 to 15 days).

Electroporation of TS Cells If a single copy of a transgene is required, such as for LoxP-flanked constructs, then electroporation is the most likely method to produce this result. However, this is the least efficient method for introducing DNA into TS cells.

ALTERNATE PROTOCOL 3

Materials Appropriate restriction enzyme Linear plasmid DNA (with mammalian antibiotic resistance gene) 3 M sodium acetate (see recipe) 70% and 100% ethanol Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) TS cells 70CM + F4H medium (see recipe) 0.05% (w/v) trypsin/1mM EDTA (see recipe) TS medium (see recipe) Microcentrifuge at 4o C Tissue culture hood 50-ml tubes 15-ml tubes (optional) Gene Pulser cuvette, 0.4 cm (Bio-Rad, cat no. 1652088) Gene Pulser electroporation device (Bio-Rad) Capacitance Extender (Bio-Rad) 100-mm culture dish Additional reagents and equipment for counting viable cells (UNIT 1C.3)

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Prepare DNA 1. Use an appropriate restriction enzyme to cut your plasmid (20 μg) of interest. Check by gel electrophoresis that digestion is complete before proceeding to the next step.

2. Heat-inactivate the enzyme (if possible) and increase the volume of the restriction digest to at least 200 μl with dH2 O. 3. Add 1/10 vol of 3 M sodium acetate (e.g., 20 μl) and then 2.5 vol of 100% ethanol (e.g., 550 μl) to precipitate DNA. A brief (∼1 hr) incubation at −20◦ C may be performed, if desired.

4. Using a microcentrifuge, spin sample 15 min at 14,000 × g, 4◦ C. Discard supernatant and wash pellet with 500 μl 70% ethanol and centrifuge again 10 min at 14,000 × g, 4◦ C. 5. Discard supernatant and allow pellet to dry briefly in a tissue culture hood. 6. Resuspend pellet in 20 μl of sterile CMF-PBS.

Prepare TS cells 7. Culture TS cells without MEFs in 70CM + F4H to ∼80% to 90% confluency. Approximately one 100-mm dish of TS cells is required per electroporation.

8. Aspirate medium and rinse twice, each time with 10 ml CMF-PBS per 100-mm dish. 9. Add 1 ml 0.05% trypsin/1 mM EDTA per dish and incubate 3 min at 37◦ C. 10. Add 9 ml TS medium per dish and break up cell aggregates by gently pipetting. 11. Transfer cell suspension into 50-ml tubes and count a 25-μl aliquot of cells with a hemacytometer (UNIT 1C.3). 12. Transfer 5 × 106 cells (or less) into a new 50-ml tube (or 15-ml tube) and centrifuge 3 min at 200 × g. 13. Aspirate supernatant and resuspend the cell pellet in 0.8 ml CMF-PBS.

Electroporate TS cells 14. Transfer 5 × 106 TS cells (in 0.8 ml CMF-PBS) into a 0.4-cm Gene Pulser cuvette. 15. Add linearized DNA (prepared in step 6) directly to the TS cell suspension in the cuvette. 16. Insert cuvette into the Gene Pulser electroporation device. 17. Set the voltage to 0.25 kV and the capacitance to 500 μFD (using the Capacitance Extender). 18. Electroporate the cells by pressing and holding the two red buttons and take note of the time constant reading from the device. A reading between 5 to 8 msec indicates the solution in the cuvette was prepared well. A time constant reading outside of this range indicates a poorly prepared sample and a new sample should be prepared if sufficient cells are available.

19. Place the cuvette with electroporated cells on ice for 20 min.

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20. Remove the TS cells from the cuvette and plate in 10 ml 70CM + F4H medium in a 100-mm culture dish. 21. Start drug-selection on the second or third day after electroporation. 22. Change the medium (70CM + F4H + drug) every second or third day until individual colonies appear (7 to 15 days). Current Protocols in Stem Cell Biology

ESTABLISHING STABLE TS LINES Once TS cells have been transfected, nucleofected, or electroporated, stably transformed clonal cell lines may be established by picking TS cell colonies in a manner quite similar to picking ES cell colonies.

SUPPORT PROTOCOL 6

Materials 70CM + 1.5× F4H medium (see recipe) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) Antibiotic (e.g., neomycin, puromycin, and zeocin) Culture of transfected TS cells (Basic Protocol 3, Alternate Protocol 2, or Alternate Protocol 3) 0.05% (w/v) trypsin/1 mM EDTA (see recipe) 4-well plates 96-well plate 20-μl adjustable pipet with appropriate tips Dissecting microscope Multichannel pipettor 1.5-ml microcentrifuge tubes 30-mm dish or 4-well plate 100-mm dish Pick TS colonies 1. Prepare sufficient 4-well plates with 250 μl 70CM + 1.5× F4H per well for the number of colonies to be picked. After 7 to 15 days of selection, ∼20 to 30 colonies/100-mm dish will appear from the electroporation protocol. More colonies are expected from the nucleofection and Lipofectamine protocols (Fig. 1E.4.3C,E,F).

2. Add 10 μl CMF-PBS to a sufficient number of wells of a 96-well plate. 3. Prepare sufficient 70CM + 1.5× F4H medium with antibiotic to have at least 300 μl per well (i.e., per colony to be picked). 4. Discard medium from dish with TS cell colonies and add 15 ml CMF-PBS. 5. Pick a TS colony together with ∼5 to 10 μl CMF-PBS using a 20-μl adjustable pipet with a the appropriate tip under a dissecting microscope. 6. Transfer the colony to a well of the 96-well plate containing 10 μl CMF-PBS (see step 2). 7. After picking 10 to 30 colonies, add 50 μl 0.05% trypsin/1 mM EDTA to each occupied well of the 96-well plate and incubate 3 to 5 min at 37◦ C. 8. Pipet gently 10 to 20 times with a multichannel pipettor to break up the colonies. 9. Add 100 μl 70CM + 1.5× F4H medium with antibiotic and remove disaggregated cells and transfer to prepared 4-well plate (step 1). 10. Add an additional 100 μl 70CM + 1.5× F4H medium with antibiotic to the same well in the 96-well plate again to collect any remaining cells and transfer to the same well of the 4-well plate. 11. Incubate 24 hr at 37◦ C. 12. Replace medium with 0.5 ml 70CM + 1.5× F4H medium with antibiotic per well. 13. Incubate at 37◦ C and change medium every 2 days. After 3 to 5 days from picking, a few colonies will grow up.

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Passage stable TS lines 14. When colonies grow and spread (5 to 7 days from picking), wash twice, each time with 500 μl CMF-PBS and add 100 μl 0.05% trypsin/1 mM EDTA and incubate 3 to 5 min at 37◦ C. 15. Add 500 μl 70CM + 1.5× F4H medium with antibiotic and pipet gently to break up colonies. 16. Transfer cells to a 1.5-ml microcentrifuge tube and wash the well with 500 μl CMF-PBS and transfer to the same 1.5-ml microcentrifuge tube. 17. Centrifuge 3 min at 200 × g. 18. Discard supernatant and resuspend cells gently in 500 μl 70CM + 1.5× F4H medium with antibiotic and seed to new 4-well plate. 19. After 5 to 10 days of feeding every other day, passage to 35-mm dish or 4-well plate again. 20. Gradually expand up to a 100-mm dish and make frozen stocks (Support Protocol 4) or use for functional analysis. BASIC PROTOCOL 4

GENERATION OF TS CELL CHIMERAS The most stringent test to determine the developmental potency of cells is the production of chimeras. ES cells are routinely used to make embryonic and adult chimeras. The two most common methods are (1) aggregation of cells to morulae and (2) microinjection of cells into the blastocoel of blastocysts. The aggregation method is not efficient with TS cells, but microinjection can give up to 20% chimeric embryos. In contrast to ES cells, TS cells and their derivatives are never found in the embryo proper, but exclusively colonize trophoblast lineages (Fig. 1E.4.4). A skilled operator that is trained in microinjection of cells into blastocysts is essential for this protocol.

Materials Genetically labeled TS cells (e.g., GFP, LacZ) 70CM + F4H medium (see recipe) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen) 0.05% (w/v) trypsin/1 mM EDTA (see recipe) TS medium (see recipe) Blastocysts (E3.5) Pseudo-pregnant females (E2.5) 14-ml round-bottom tubes (BD Falcon) Microinjection facility with operator Dissecting microscope with UV fluorescence Prepare TS cells for microinjection 1. Culture genetically labeled TS cells without MEFs in 70CM + F4H to ∼80% to 90% confluency. One 60-mm dish of TS cells is sufficient for microinjection.

Isolation and Manipulation of Mouse Trophoblast Stem Cells

Before using TS cells to generate chimeras they should exhibit ∼10% (or less) differentiation. If the levels of differentiation appear higher than this, the cells can be differentially plated to enrich for stem cells (see Support Protocol 3).

2. Aspirate medium and rinse twice, each time with 5 ml CMF-PBS per 60-mm dish. 3. Add 0.5 ml 0.05% trypsin/1 mM EDTA per dish and incubate 3 min at 37◦ C.

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Figure 1E.4.4 TS cell chimera. An 8.5 dpc TS cell chimera generated by injecting GFP-labeled TS cells into a blastocyst. The embryo was observed under partial bright-field and UV optics (A) and dark-field optics (B). (C) A sketch of the conceptus indicates the embryo (e), decidua (d), and placenta (p).

4. Add 4.5 ml TS medium per dish and break cell aggregates by gently pipetting to generate a single-cell suspension. 5. Transfer to a round-bottom tube, place on ice, and bring to microinjection operator.

Inject blastocyst with TS cells 6. Using the microinjection operator, inject TS cells into E3.5 blastocysts using techniques identical to those used to inject ES cells (Nagy et al., 2003). Inject 5 to 10 TS cells per blastocyst and use up to 60 blastocysts. Smaller cells should be chosen for injection, since the size of TS cells increase as they differentiate and TS cell cultures are invariably heterogeneous.

7. Transfer up to twelve injected blastocysts per E2.5 pseudo-pregnant females.

Analyze chimeras 8. Dissect embryos from E5.5 to just before term (E18.5). Take special care to keep the trophoblast tissue intact. If TS cells were labeled with GFP or an alternate fluorescent protein, chimeras may be identified using a fluorescence dissecting microscope (see Fig. 1E.4.4). Chimeras may be stored at 4◦ C in CMF-PBS with azide for a short time (1 to 2 weeks) or fixed in 4% paraformaldehyde for long-term storage at 4◦ C. Fixation may reduce the fluorescence of GFP.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

BSA, 0.1% (w/v)/CMF-PBS Dissolve 11 mg fraction V bovine serum albumin (BSA) in 11 ml phosphatebuffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen). Filter through a 0.45-μm filter. Store in 1.05-ml aliquots up to several years at −80◦ C.

DMEM/10% (v/v) FBS Dulbecco’s modified Eagle medium (DMEM; pH 7.2) supplemented with: 10% (v/v) FBS 50 U/ml penicillin and 50 μg/ml streptomycin, optional Store up to 2 months at 4◦ C

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EDTA (1 mM)/CMF-PBS Dissolve 0.19 g of EDTA·4Na in 500 ml phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Invitrogen). Sterilize by filtration or autoclave. Store up to 6 months at 4◦ C.

Freezing medium for MEFs, 2× 50% FBS, 20% dimethyl sulfoxide (DMSO) in DMEM/10% FBS. Add 5 ml FBS and 2 ml DMSO to 3 ml DMEM/10% FBS (see recipe). Prepare fresh before use.

Freezing medium for TS cells, 2× 50% FBS, 20% dimethyl sulfoxide (DMSO) in TS medium. Add 5 ml FBS and 2 ml DMSO to 3 ml TS medium (see recipe). Prepare fresh before use.

Heparin, 1 mg/ml (1000× stock) Resuspend heparin (Sigma, cat no. H3149) in sterile CMF-PBS (Invitrogen) to a concentration of 1.0 mg/ml and store in 100-μl aliquots up to several years at −80◦ C. Thaw aliquots as needed and store up to 3 months at 4◦ C; do not refreeze.

Human recombinant FGF4, 25 μg/ml (1000× stock) Add 1 ml 0.1% (w/v) BSA/CMF-PBS (see recipe) directly to a vial of lyophilized human recombinant FGF4 (25 μg; PeproTech, cat. no. 100-31). Mix well by gentle pipetting and freeze in 100-μl aliquots up to several years at −80◦ C. Thaw aliquots as needed and store up to 1 month at 4◦ C; do not refreeze. Filter sterilization is not necessary, since the BSA/CMF-PBS solution is already sterile. Recombinant FGF1 (aFGF) and FGF2 (bFGF) have also been successfully used in this protocol, and they are slightly cheaper than FGF4.

Mitomycin C (MMC) Wearing protective gloves, flip off the plastic button top of a vial containing 2 mg MMC (Sigma-Aldrich, cat no. M0503) and inject 2 ml sterile water into the vial. Store this 1 mg/ml stock solution up to 1 week at 4◦ C in the dark according to the manufacturer’s data sheet. CAUTION: Mitomycin C is VERY TOXIC. Further filtration is not usually required. We have, however, empirically found that the stock solution can be kept frozen in aliquots at –20◦ C for at least 1 year without any noticeable decrease in its activity (do not refreeze once thawed).

70CM + F4H medium TS medium containing 70% MEF-conditioned medium and 25 ng/ml FGF4 and 1 μg/ml heparin. Add 10 μl each of FGF4 (see recipe) and heparin stock solutions (see recipe) to 3 ml TS medium (see recipe) and 7 ml MEF-conditioned medium (MEF-CM; see Support Protocol 2). Prepare fresh before use.

70CM + 1.5× F4H medium

Isolation and Manipulation of Mouse Trophoblast Stem Cells

TS medium containing 70% MEF-CM and 37.5 ng/ml FGF4 and 1.5 μg/ml heparin. Add 15 μl each of FGF4 (see recipe) and heparin stock solutions (see recipe) to 3 ml TS medium (see recipe) and 7 ml MEF-conditioned medium (MEF-CM; see Support Protocol 2). Prepare fresh before use.

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Sodium acetate, 3M Add 123.05 g of sodium acetate (anhydrous) to 400 ml dH2 O. Adjust the pH to 7.0 with dilute acetic acid. Adjust volume to 500 ml with dH2 O and autoclave. Store up to several years at room temperature.

Trypsin (0.1%)/EDTA (1 mM) Dilute 0.25% trypsin/1 mM EDTA·4Na (Invitrogen) with 1.5× vol of 1 mM EDTA/CMF-PBS (see recipe). Store up to 2 months at 4◦ C.

Trypsin (0.05%)/EDTA (1 mM) Dilute 0.25% trypsin/1 mM EDTA·4Na (Invitrogen) with 4× vol of 1 mM EDTA/CMF-PBS (see recipe). Store up to 2 months at 4◦ C.

Trypsin (0.5%)/pancreatin (2.5%)/EDTA (1 mM) Mix 8 ml PBS, 2 ml 2.5% trypsin (Invitrogen), and 20 μl 0.5 M EDTA in a 15-ml tube. Add 2.5 g pancreatin powder (any brand) and gently mix by inverting the tube for ∼5 min at room temperature. Incubate an additional 10 to 20 min on a rotator a 4◦ C. Divide into 1-ml aliquots and store up to 1 year at −20◦ C. Do not refreeze aliquots once thawed. This solution will not become clear and some debris will remain undissolved.

TS medium RPMI 1640 (Invitrogen ) supplemented with: 20% (v/v) fetal bovine serum (FBS; any brand; batch-tested for ES cells, if possible) 2 mM L-glutamine 1 mM sodium pyruvate 100 mM 2-mercaptoethanol 50 U/ml penicillin and 50 μg/ml streptomycin Store up to 1 month at 4◦ C Penicillin and streptomycin can be omitted if preparing medium for lipofection (Basic Protocol 3).

TS + F4H medium TS medium containing 25 ng/ml FGF4 and 1 μg/ml heparin. Add 10 μl each of FGF4 (see recipe) and heparin (1000× stock solutions; see recipe) to 10 ml TS medium (see recipe). Prepare fresh prior to each use.

TS + 1.5× F4H medium TS medium containing 37.5 ng/ml FGF4 and 1.5 μg/ml heparin. Add 15 μl each of FGF4 (25 μg/ml; see recipe) and heparin (1 mg/ml; see recipe) stock solutions to 10 ml TS medium (see recipe). Prepare fresh prior to each use.

COMMENTARY Background Information During early mouse development the first segregation of cell lineages occurs at 3.5 days post-coitum (3.5 dpc). At this stage, the newly formed blastocyst is composed of only two cell types—the inner cell mass (ICM) and trophectoderm (TE). Within the next day a third lineage is formed and becomes apparent on the surface of the ICM, the primitive

endoderm. Stable, permanent stem cell lines have been derived from each of these early embryonic lineages. The most well-known, embryonic stem (ES) cells, are derived from the ICM or early epiblast (4.5 dpc) in the presence of feeder cells, which provide the critical cytokine Leukemia inhibitory factor (LIF; Evans and Kaufman, 1981; Martin, 1981; Smith et al., 1988; Brook and Gardner, 1997).

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In contrast, the primitive endoderm can give rise to extraembryonic endoderm (XEN) cell lines without the addition of exogenous cytokines (Kunath et al., 2005). The derivation of mouse trophoblast stem (TS) cells also requires a feeder layer of mouse embryonic fibroblasts (MEFs), plus the critical ligand FGF4 (Tanaka et al., 1998). TS cells have also been successfully isolated from the extraembryonic ectoderm (ExE) of 6.5 dpc embryos and the chorionic ectoderm (ChE) of 7.5 dpc embryos (Tanaka et al., 1998; Uy et al., 2002). The critical signaling molecule, FGF4, has been implicated in maintenance of trophoblast progenitors from a number of gene targeting and gene expression studies. The expression of Fgf4 in the ICM and early epiblast (Niswander and Martin, 1992) and reciprocal expression of Fgfr2 in the overlying ExE (Ciruna and Rossant, 1999; HaffnerKrausz et al., 1999), suggested that paracrine FGF signaling from the epiblast is important to maintain the early trophoblast lineage in vivo. This model was further supported by evidence that activation of the MAPKs Erk1/2 in the ExE is FGF-dependent (Corson et al., 2003). Embryos mutant for Fgf4 or Fgfr2 die shortly after implantation and do not exhibit any trophoblast expansion (Feldman et al., 1995; Arman et al., 1998). Some downstream components of this pathway, such as Grb2, FRS2α, and Erk2, exhibit similar trophoblast defects when mutated (Cheng et al., 1998; Saba-ElLeil et al., 2003; Gotoh et al., 2005). A second signaling molecule(s), distinct from LIF, was suggested by the need for mouse embryonic fibroblasts (MEFs) or MEF-conditioned medium (MEF-CM) to maintain TS cells in culture. Investigations by Erlebacher and colleagues identified the active components in MEF-CM to be TGFβ and the related ligand activin (Erlebacher et al., 2004). They were able to maintain and derive TS cell lines with recombinant TGFβ or activin in the absence of MEFs or MEF-CM. The molecule in vivo that activates this pathway (Smad2 and Samd3) may be maternally derived activin or epiblastderived Nodal (a TGFβ-related ligand). Both are expressed at the right time and Nodal null embryos exhibit trophoblast defects by 9.5 dpc (Albano et al., 1994; Ma et al., 2001). TS cells can also be directly derived from ES cells through manipulation of lineagedeterminant transcription factors. Oct4 is a critical transcription factor for the ICM and ES cells (Nichols et al., 1998). Repression of this gene in ES cells caused trophoblast differ-

entiation, and stable TS cell lines can be derived if FGF4 is supplied to the culture when Oct4 is down-regulated (Niwa et al., 2000). The caudal-related protein Cdx2 and the T-box transcription factor Eomesodermin (Eomes) are critically important for early trophoblast development (Russ et al., 2000; Strumpf et al., 2005). Over-expression of either Cdx2 or Eomes in ES cells results in differentiation to TS cells in the appropriate culture conditions (Niwa et al., 2005). More recently, two methods have been described to derive TS cells from ES cells without genetic manipulation. In the first method, collagen IV plates are used in combination with TS cell medium. Interestingly, TS cell lines could only be derived from feeder-dependent ES cell lines (SchenkeLayland et al., 2007). In a second study, Wnt3a was found to induce Cdx2 expression in ES cells. Combining Wnt3a and LIF-removal resulted in the highest Cdx2 induction with subsequent establishment of TS cell-like cultures (He et al., 2008).

Critical Parameters and Troubleshooting Unlike ES cells, TS cells attach directly to the bottom of tissue culture plates and push MEFs aside as they expand, rather than growing on top of MEFs. Half the number of MEFs, compared to ES cell coculture, are therefore used with TS cells to leave space for colony expansion. The presence of too many MEFs may cause physical stress, which seems to induce spontaneous differentiation of TS cells. If TS cells, cocultured with MMC-MEFs, are being passaged at a high dilution (e.g., 1 in 20) to new MMC-MEFs, the removal of the older MMC-MEFs is not necessary. However, if they are to be passed at a low dilution (e.g., 1 in 5), removal of old MMC-MEFs from the mixed suspension is recommended (see Support Protocol 3). Low efficiency in a derivation of TS cell lines and unexpected differentiation of TS cells during maintenance are occasionally caused by low quality of MEF/MEF-CM. It is recommended to verify the ability of new batches of MMC-MEF and MEF-CM to maintain already established TS cell lines. Note that the appropriate number of MMC-MEFs for coculture with TS cells and to prepare MEF-CM described in this unit are based on cell counts before freezing of MMC-treated MEFs. Therefore, the actual number of viable MEFs after thawing a frozen stock should be less than those shown in Tables 1E.4.1 and

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1E.4.2 and may vary depending on freezing conditions. If small numbers of MMC-MEFs appear to survive after thawing frozen stocks, simply increasing the number of MMC-MEFs to inoculate may sometimes solve this problem. Mitomycin C treatment of MEFs just before use (i.e., without freezing) is another option to consider. It is very difficult, if not impossible, to completely block spontaneous differentiation of TS cells even in the presence of increased amounts of FGF4. Many of the differentiated cells are trophoblast giant cells. However, it is fortuitous that giant cells are quite resistant to trypsin treatment, which results in a partial enrichment of true TS cells at each passage. Differential plating of cells (Support Protocol 3) can also be used to reduce the amount of differentiated cells during a passage. During the early passages of establishing new TS cell lines, the culture can sometimes appear to be entirely differentiated, especially into giant cells. This can also occur at later passages, if one of the reagents is off. Do not despair if this occurs. Although the culture may appear to have lost all TS cell colonies, we recommend that you continue feeding the culture for up to 15 days without passaging. In most cases, TS cell colonies begin to appear, seemingly out of nowhere. The presence of a large number of giant cells does not inhibit the emergence or growth of TS cell colonies. If these cultures are on MMC-MEFs, they may be supplemented with MEF-CM once the MMC-MEFs appear to be dying. During the first 5 to 10 passages it is difficult to decide when the cells are ready to be passaged, especially since the act of passaging with trypsin seems to cause differentiation during these early stages. It is best to wait until almost half of the well is covered with true TS cell colonies (i.e., not giant cells), or until the individual colonies appear overgrown. It is not uncommon to go 10 days between passages. If there is doubt about whether to passage or not, then we recommend you simply feed the culture and re-assess the next day.

Anticipated Results The derivation of TS cell lines from blastocysts is highly efficient when permissive mouse strains are used (e.g., ICR [CD-1], 129/sv and 129 substrains). For example, 58 clonal TS cell lines were established from 91 blastocysts (64%) and 17 from 39 6.5 dpc embryos (44%), respectively, of 129/Sv and ICR mice (Tanaka et al., 1998). The efficiency slightly declines when C57BL/6 (BL6) back-

ground is introduced. For example, 5 lines were established from 10 blastocysts (50%) of BL6/129 mixed background mouse (Arima et al., 2006). Since the original publication the efficiency of derivation from permissive strains is ∼80% from blastocysts and ∼90% from 6.5 dpc ExE or 7.5 dpc ChE is ∼90%. There is much less contamination of XEN cells when TS cells are derived from postimplantation embryos (i.e., ExE and ChE). For either procedure expect a large amount of differentiation for the first 10 passages. Once the TS cell lines are well established, differentiated cells still appear at frequencies of up to 10%. Introducing DNA into TS cells has been a challenge. The modified Lipofectamine protocol described in Basic Protocol 3 is the most efficient with 20% to 30% of the cells transfected. The nucleofection and electroporation protocols are suitable for deriving stably transformed TS cell lines with potential single-site integrations. However, only 20 to 50 colonies are obtained, in contrast to hundreds of colonies for similar procedures performed with ES cells. Due to this low efficiency, gene targeting is not recommended in TS cells. If genetically null TS cells are desired, it is recommended to perform the targeting in ES cells first. Then generate chimeric mice to get heterozygous mice from which homozygous null embryos are obtained to be used for TS cell derivation as described in Basic Protocol 1 and Alternate Protocol 1. This has been successfully performed for several genes, including Arnt, Ink4a, and Dnmt3l (Adelman et al., 2000; Erlebacher et al., 2002; Arima et al., 2006). Alternatively, both alleles may be targeted and the resulting null ES cells can be directly differentiated into TS cells by overexpression of Cdx2 or by using culture conditions that induce TS cells (Niwa et al., 2005; Schenke-Layland et al., 2007; He et al., 2008). The generation of TS cell chimeras is not trivial. In ideal conditions, expect 20% of recovered embryos to be chimeric. However, high-contribution chimeras, where more than half of the trophoblast tissue is derived from injected TS cells, are found in <5% of embryos. As recommended in Basic Protocol 4, differential plating to enrich for true TS cells will help increase the number of chimeras. If facilities exist, fluorescence-activated cell sorting of live cells for 2N DNA content with Hoechst 33342 (Karawajew et al., 1990) significantly reduces the heterogeneity of TS cells for several passages and improves chimera formation.

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Time Considerations Before attempting to derive TS cell lines from embryos, sufficient stocks of frozen MMC-MEFs (Support Protocol 1) and MEFconditioned medium (Support Protocol 2) need to be generated. If MEFs are already available, these stocks will require 2 weeks to make. If MEFs are to be derived from embryos, an additional 3 weeks is required. Once all the reagents are prepared, matings are needed to get 3.5 dpc or 6.5 dpc pregnant females. Isolation of blastocysts and placing into TS cell conditions requires ∼1 to 2 hr, while isolation of ExE from 6.5 dpc embryos can take ∼3 to 4 hr depending on the embryo dissection skills of the experimenter. The subsequent steps in the TS cell derivation protocol required up to 1 hr every second day. The early passages of TS cells grow very slowly and it can be up to 10 days between passages. There is also significant differentiation for the first 7 to 10 passages. It can take up to 3 months to derive a TS cell line that can be consistently passaged and used for experiments, such as transfection and chimera formation.

Acknowledgements We thank Adrian Erlebacher for sharing unpublished information regarding the culture conditions for TS cells and Keisuke Kaji for the nucleofection protocol.

Literature Cited Adelman, D.M., Gertsenstein, M., Nagy, A., Simon, M.C., and Maltepe, E. 2000. Placental cell fates are regulated in vivo by HIF-mediated hypoxia responses. Genes Dev. 14:3191-3203. Albano, R.M., Arkell, R., Beddington, R.S., and Smith, J.C. 1994. Expression of inhibin subunits and follistatin during postimplantation mouse development: Decidual expression of activin and expression of follistatin in primitive streak, somites and hindbrain. Development 120:803813. Arima, T., Hata, K., Tanaka, S., Kusumi, M., Li, E., Kato, K., Shiota, K., Sasaki, H., and Wake, N. 2006. Loss of the maternal imprint in Dnmt3Lmat−/− mice leads to a differentiation defect in the extraembryonic tissue. Dev. Biol. 297:361-373. Arman, E., Haffner-Krausz, R., Chen, Y., Heath, J.K., and Lonai, P. 1998. Targeted disruption of fibroblast growth factor (FGF) receptor 2 suggests a role for FGF signaling in pregastrulation mammalian development. Proc. Natl. Acad. Sci. U.S.A. 95:5082-5087. Isolation and Manipulation of Mouse Trophoblast Stem Cells

Brook, F.A. and Gardner, R.L. 1997. The origin and efficient derivation of embryonic stem cells in the mouse. Proc. Natl. Acad. Sci. U.S.A. 94:5709-5712.

Cheng, A.M., Saxton, T.M., Sakai, R., Kulkarni, S., Mbamalu, G., Vogel, W., Tortorice, C.G., Cardiff, R.D., Cross, J.C., Muller, W.J., and Pawson, T. 1998. Mammalian Grb2 regulates multiple steps in embryonic development and malignant transformation. Cell 95:793-803. Ciruna, B.G. and Rossant, J. 1999. Expression of the T-box gene Eomesodermin during early mouse development. Mech. Dev. 81:199-203. Corson, L.B., Yamanaka, Y., Lai, K.-M.V., and Rossant, J. 2003. Spatial and temporal patterns of ERK signaling during mouse embryogenesis. Development 130:4527-4537. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Erlebacher, A., Lukens, A.K., and Glimcher, L.H. 2002. Intrinsic susceptibility of mouse trophoblasts to natural killer cell-mediated attack in vivo. Proc. Natl. Acad. Sci. U.S.A. 99:1694016945. Erlebacher, A., Price, K.A., and Glimcher, L.H. 2004. Maintenance of mouse trophoblast stem cell proliferation by TGF-beta/activin. Dev. Biol. 275:158-169. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Feldman, B., Poueymirou, W., Papaioannou, V.E., DeChiara, T.M., and Goldfarb, M. 1995. Requirement of FGF-4 for postimplantation mouse development. Science 267:246-249. Gardner, R.L. and Davies, T.J. 2000. Mouse chimeras and the analysis of development. Methods Mol. Biol. 135:397-424. Gotoh, N., Manova, K., Tanaka, S., Murohashi, M., Hadari, Y., Lee, A., Hamada, Y., Hiroe, T., Ito, M., Kurihara, T., Nakazato, H., Shibuva, M., Lax, I., Lacy, E., and Schlessinger, J. 2005. The docking protein FRS2alpha is an essential component of multiple fibroblast growth factor responses during early mouse development. Mol. Cell Biol. 25:4105-4116. Haffner-Krausz, R., Gorivodsky, M., Chen, Y., and Lonai, P. 1999. Expression of Fgfr2 in the early mouse embryo indicates its involvement in preimplantation development. Mech. Dev. 85:167-172. He, S., Pant, D., Schiffmacher, A., Meece, A., and Keefer, C.L. 2008. Lymphoid enhancer factor 1mediated wnt signaling promotes the initiation of trophoblast lineage differentiation in mouse embryonic stem cells. Stem Cells 26:842-849. Karawajew, L., Rudchenko, S., Wlasik, T., Trakht, I., and Rakitskaya, V. 1990. Flow sorting of hybrid hybridomas using the DNA stain Hoechst 33342. J. Immunol. Methods 129:277-282. Kunath, T., Arnaud, D., Uy, G.D., Okamoto, I., Chureau, C., Yamanaka, Y., Heard, E., Gardner, R.L., Avner, P., and Rossant, J. 2005. Imprinted X-inactivation in extra-embryonic endoderm cell lines from mouse blastocysts. Development 132:1649-1661.

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Lakshmipathy, U., Buckley, S., and Verfaillie, C. 2007. Gene transfer via nucleofection into adult and embryonic stem cells. Methods Mol. Biol. 407:115-126. Ma, G.T., Soloveva, V., Tzeng, S.J., Lowe, L.A., Pfendler, K.C., Iannaccone, P.M., Kuehn, M.R., and Linzer, D.I. 2001. Nodal regulates trophoblast differentiation and placental development. Dev. Biol. 236:124-135.

determines trophectoderm differentiation. Cell 123:917-929. Russ, A.P., Wattler, S., Colledge, W.H., Aparicio, S.A., Carlton, M.B., Pearce, J.J., Barton, S.C., Surani, M.A., Ryan, K., Nehls, M.C., Wilson, V., and Evans, M.J. 2000. Eomesodermin is required for mouse trophoblast development and mesoderm formation. Nature 404:95-99.

Mak, W., Baxter, J., Silva, J., Newall, A.E., Otte, A.P., and Brockdorff, N. 2002. Mitotically stable association of polycomb group proteins eed and enx1 with the inactive x chromosome in trophoblast stem cells. Curr. Biol. 12:1016-1020.

Saba-El-Leil, M.K., Vella, F.D., Vernay, B., Voisin, L., Chen, L., Labrecque, N., Ang, S.L., and Meloche, S. 2003. An essential function of the mitogen-activated protein kinase Erk2 in mouse trophoblast development. EMBO Rep. 4:964968.

Martin, G.R. 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. 78:76347638.

Schenke-Layland, K., Angelis, E., Rhodes, K.E., Heydarkhan-Hagvall, S., Mikkola, H.K., and Maclellan, W.R. 2007. Collagen IV induces trophoectoderm differentiation of mouse embryonic stem cells. Stem Cells 25:1529-1538.

Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. 2003. Manipulating the mouse embryo: A laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.

Smith, A.G., Heath, J.K., Donaldson, D.D., Wong, G.G., Moreau, J., Stahl, M., and Rogers, D. 1988. Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336:688-690.

Nichols, J., Zevnik, B., Anastassiadis, K., Niwa, H., Klewe-Nebenius, D., Chambers, I., Scholer, H., and Smith, A. 1998. Formation of pluripotent stem cells in the mammalian embryo depends on the POU transcription factor Oct4. Cell 95:379391. Niswander, L. and Martin, G.R. 1992. Fgf-4 expression during gastrulation, myogenesis, limb and tooth development in the mouse. Development 114:755-768. Niwa, H., Miyazaki, J., and Smith, A.G. 2000. Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat. Genet. 24:372-376. Niwa, H., Toyooka, Y., Shimosato, D., Strumpf, D., Takahashi, K., Yagi, R., and Rossant, J. 2005. Interaction between Oct3/4 and Cdx2

Strumpf, D., Mao, C.A., Yamanaka, Y., Ralston, A., Chawengsaksophak, K., Beck, F., and Rossant, J. 2005. Cdx2 is required for correct cell fate specification and differentiation of trophectoderm in the mouse blastocyst. Development 132:2093-2102. Tanaka, S., Kunath, T., Hadjantonakis, A.K., Nagy, A., and Rossant, J. 1998. Promotion of trophoblast stem cell proliferation by FGF4. Science 282:2072-2075. Uy, G.D., Downs, K.M., and Gardner, R.L. 2002. Inhibition of trophoblast stem cell potential in chorionic ectoderm coincides with occlusion of the ectoplacental cavity in the mouse. Development 129:3913-3924.

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Isolation of Amniotic Mesenchymal Stem Cells

UNIT 1E.5

Fabio Marongiu,1 Roberto Gramignoli,1 Qian Sun,1 Veysel Tahan,1 Toshio Miki,1 Kenneth Dorko,1 Ewa Ellis,1 and Stephen C. Strom1 1

University of Pittsburgh, Pittsburgh, Pennsylvania

ABSTRACT Mesenchymal stem cells (MSCs) have the ability to differentiate into osteocytes, chondrocytes, and adipocytes and possess immunomodulatory properties. Amniotic membrane from human term placenta is a potential source of multipotent MSCs that could be useful for regenerative medicine. This unit describes a detailed and simple protocol for the isolation of amniotic mesenchymal cells. We also introduce a simple density separation technique for the purification of this cell type from possible contamination with amniotic C 2010 by John Wiley epithelial cells. Curr. Protoc. Stem Cell Biol. 12:1E.5.1-1E.5.11.  & Sons, Inc. Keywords: placenta r amnion r mesenchymal stem cells r MSC

INTRODUCTION The amnion is a thin, avascular membrane composed of a compact layer of cuboidal and columnar epithelial cells, which are in contact with the amniotic fluid on the external side and attached to a basal lamina on the inner side. This lamina is connected to the amniotic mesoderm, a layer of extracellular components (collagens and fibronectin) into which is embedded a network of dispersed fibroblast-like mesenchymal cells (Fig. 1E.5.1). During the development of the embryo, the placental tissue originates before gastrulation, supporting the hypothesis that some cells from this tissue may retain multipotent/pluripotent characteristics. Two types of stem cells can be isolated from the amnion of human term placentas: amniotic epithelial cells (hAECs) and amniotic mesenchymal cells (hAMSCs). The amniotic epithelium develops from the epiblast by 8 days after fertilization and prior to gastrulation. It has been reported that these cells, isolated from human term placenta, express surface makers and transcription factors normally present on embryonic stem and germ cells, and these cells have the ability to differentiate into cells from all three germ layers (Miki et al., 2005). hAECs do not express telomerase, do display a normal karyotype, and are nontumorigenic upon transplantation (Miki and Strom, 2006). A detailed protocol for the isolation of hAECs has been published (UNIT 1E.3). Mesenchymal stem cells (MSC) are multipotent cells that differentiate into osteoblasts, myocytes, chondrocytes, and adipocytes. They have been isolated from a variety of tissues, including bone marrow, adipose tissue, Wharton’s jelly, umbilical cord blood, and different compartments of placenta (Parolini et al., 2008). Human amnion-derived mesenchymal stem cells (hAMSCs) have fibroblast-like morphology, have the ability to form colonies, and can be subcultured several times (Soncini et al., 2007). Recent studies demonstrated that AM cells have immunomodulatory properties and can strongly inhibit T lymphocyte proliferation (Magatti et al., 2008). The availability of tissue and the absence of ethical concerns for the tissue source make placenta a potentially useful source of stem cells for cell transplantation and regenerative medicine. Current Protocols in Stem Cell Biology 1E.5.1-1E.5.11 Published online March 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808 C 2010 John Wiley & Sons, Inc. Copyright 

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epithelial cells

mesenchymal cells

Figure 1E.5.1

Cross-section of human amnion.

In this protocol, we describe a method to isolate hAMSCs following the isolation of amniotic epithelial cells (UNIT 1E.3) from term placentas obtained from cesarean section procedures (Basic Protocol). Furthermore, we introduce a method to purify hAMSC from possible epithelial cell contamination by a simple Percoll gradient separation technique (Alternate Protocol). NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: This protocol uses human tissues. It should be submitted to, reviewed, and approved by the appropriate institutional review board, and all tissues should be obtained with the informed consent of the source. BASIC PROTOCOL

Isolation of Amniotic Mesenchymal Stem Cells

ISOLATION OF HUMAN AMNIOTIC MESENCHYMAL STEM CELLS Human amniotic mesenchymal stem cells (hAMSCs) are isolated from human amnion that, like other placental tissues, would normally be discarded post delivery. For sterility purposes, placentas are normally obtained from cesarean section; however, theoretically, all placentas should be useful for cell isolation. Tissues are obtained with local Institutional Review Board (IRB) approval in the U.S., or under appropriate Ethical Committee approval. For the studies described here, all infectious pathogen-positive deliveries including HBV, HCV, and HIV, and prediagnosed genetic abnormalities cases are excluded. Even with these precautions, all staff members are made aware that the tissue should be considered potentially infectious material and standard precautions for safe use of human tissue must be followed. If you also need to isolate amniotic epithelial cells (hAECs), please refer to UNIT 1E.3 for the detailed protocol. Then, proceed with the isolation of hAMSCs starting from step 13 of this protocol.

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Materials Hanks’ balanced salt solution (HBSS, calcium- and magnesium-free; Lonza, cat. no. 04-315Q) 70% (w/v) ethanol 0.05% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25300-054) Term placenta, freshly delivered Digestion Solution (see recipe) Standard culture medium (see recipe) Laminar flow cabinet equipped with the following: Absorbent bench paper Sterile field 16 × 29–in. Sterile scalpel 500-ml beakers (2) Sterile Scissors (2) and forceps (2) Sterile gloves and sleeves 100-μm nylon cell strainers (4) 50-ml Falcon centrifuge tubes (8) Centrifuge 37◦ C incubator 37◦ C water bath Additional reagents and equipment for cell counting (UNIT 1C.3) Prepare hood for isolation 1. Equip a laminar flow cabinet BSL-II with the following (Fig. 1E.5.2): Absorbent bench paper Sterile field 16 × 29–in. Sterile scalpel Sterilized 500-ml beaker Sterilized scissors and forceps Sterile gloves and sleeves 100-μm nylon cell strainers 50-ml centrifuge tubes. 2. Add 200 ml HBSS to a sterile beaker.

Carry out the dissection 3. Wear sterile gloves and sleeves and place the whole placenta on the sterile-field (Fig. 1E.5.3). The maternal surface (rough surface) should be facing down on the bench paper with the smooth surface with the umbilical cord facing up. In this position, the amnion membrane will lay across the upper surface of the placenta.

4. Trim the umbilical cord close to the placental surface and cut an X-shaped incision into, but not through, the placental tissue (Fig. 1E.5.4). The X should intersect at the position of the umbilical cord. It may be easiest to start the incision in the region of the umbilical cord. After this step, there could be an excess of blood on top of the amnion. If that is the case, pour some HBSS on top of the membrane and gently move it away by hand to remove blood clots. Embryonic and Extraembryonic Stem Cells

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Figure 1E.5.2

Sterile equipment for the isolation of hAECs.

Figure 1E.5.3

Placenta with amnion membrane facing up.

5. Peel the amnion membrane from the underlying chorion layer of the placenta body. Start from the cut edge (middle of the placenta body) and peel the membrane from the placenta (Fig. 1E.5.5). Isolation of Amniotic Mesenchymal Stem Cells

6. Place the amnion in the sterilized 500-ml beaker containing 200 ml HBSS from step 2 (Fig. 1E.5.6).

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Figure 1E.5.4

Placenta with an X-shaped incision.

Figure 1E.5.5

Peeling the amnion membrane.

7. Discard the remaining part of the placenta, remove the dissecting material, and wipe the hood with ethanol. 8. Wash the amnion two to three times with ∼200 ml HBSS, each time by moving the amnion to a clean beaker with sterile forceps. This washing step is crucial for the trypsin to work properly. Blood clots will reduce the efficiency of the trypsin. Ideally, all clots on the membrane will be cleared.

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Figure 1E.5.6

Washing the amniotic membrane in HBSS.

Release amniotic epithelial cells This is a simplified digestion procedure to be used when hAECs are not needed. If you also wish to isolate hAECs, please refer to UNIT 1E.3 (Miki et al., 2005) for a detailed protocol. Then, proceed with the isolation of hAMSCs starting from step 13 of this protocol. 9. Thaw and prewarm the 0.05% (w/v) trypsin/EDTA solution in a water bath to 37◦ C. 10. Place the membrane into two 50-ml centrifuge tubes and add ∼20 ml of 0.05% (w/v) trypsin/EDTA. Incubate 1 hr at 37◦ C. The volume of membrane tissue from an average placenta is ∼20 ml.

11. Vigorously shake the tubes in order to release hAECs. 12. Transfer the membrane into a 500-ml beaker with cold HBSS. Discard the trypsin digest. 13. Wash the amnion two to three times with ∼200 ml HBSS, each time by moving the amnion to a clean beaker.

Digest the membrane 14. Transfer the membrane into two (or more) 50-ml centrifuge tubes, allowing excess HBSS to drip from the membrane. The amount of wet tissue will be increased after trypsin digestion. In order not to dilute the collagenase solution excessively, divide the tissue as needed so as not to have more than 20 ml of wet tissue in 50 ml final volume.

15. Add the digestion solution to completely fill the tubes. Incubate on a rotator 1 hr at 37◦ C. Isolation of Amniotic Mesenchymal Stem Cells

The incubation time varies according to the membrane thickness, ranging between 45 min to 1.5 hr. It is wise to occasionally check the status of digestion after the first 30 min, and stop the incubation as soon as the tissue is completely dissolved.

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16. Equally divide the contents of each tube into two new 50-ml tubes. 17. Add an equal volume of cold HBSS and centrifuge 5 min at ∼200 × g, 4◦ C. 18. Discard the supernatant. Resuspend the pellet with fresh HBSS to fill the tube and centrifuge again 5 min at ∼200 × g, 4◦ C. 19. Resuspend the pellets in a small volume of standard culture medium and combine them in a single solution. 20. Count the cell number (UNIT 1C.3). 21. If cells are to be cultured, plate up to 1 × 105 cells per cm2 in standard culture medium. If cells are to be cultured, plate up to 1 × 105 cells per cm2 in standard culture medium, changing the medium twice a week. Subculture (1:4) the cells, as needed, when the cells exceed 70% confluence.

PURIFICATION OF AMNIOTIC MESENCHYMAL CELLS FROM AMNIOTIC EPITHELIAL CELLS

ALTERNATE PROTOCOL

Although the isolation of hAECs by trypsin/EDTA digestion yields a virtually pure preparation, this is not the case for hAMSC isolation; a considerable number of hAECs may still be attached to the membrane before collagenase digestion, and this may result in the presence of hAECs in the hAMSC preparation. Considering the slow adhesion of hAECs to culture substrates, as opposed to the fast attachment of hAMSCs, it is good practice to change the culture medium 1 to 2 hr after plating. This will remove many of the hAECs that might be present in the preparation. However, significant contamination of hAMCs with hAECs will frequently occur. The morphological differences between the two cell types will allow you to easily determine whether the contamination with hAECs is substantial or not after 24 hr in culture. In this protocol, we describe a density separation method to purify hAMSCs from hAECs. NOTE: The following protocol will work best with cells cultured for 24 hr, as the two cell types acquire different morphologies and cellular densities when attached to culture substrates. The procedure might be unsuccessful if performed with freshly isolated cells.

Additional Materials (also see Basic Protocol) Isolated amniotic mesenchymal cells in culture (see the Basic Protocol) 0.25% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25200-056) Percoll (Sigma, cat. no. P4937) 10× HBSS (Invitrogen, cat. no. 14065-056) Hanks’ balanced salt solution, calcium- and magnesium-free (HBSS-CMF; Lonza, cat. no. 04-3150) 15-ml conical tubes Prepare cell suspension 1. After 1 day in culture, rinse hAMSC cultures twice with HBSS-CMF to remove traces of serum. Wash the cells in HBSS twice prior to trypsin digestion in order to remove serum whose components inhibit the enzyme’s activity.

2. Release hAMSCs from culture substrate by 0.25% (w/v) trypsin/EDTA digestion (∼5 min at 37◦ C).

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3. Count the cell number (UNIT 1C.3). 4. Resuspend the cells in standard culture medium to a density of 30 × 106 cells per ml. 5. Place on ice.

Prepare Percoll gradients 6. As instructed by the manufacturer, prepare the 100% Percoll by mixing 9 parts pure Percoll with 1 part 10× HBSS. For 30 × 106 cells, ∼6 ml of 100% Percoll will be needed (5.4 ml pure Percoll + 0.6 ml 10× HBSS). All the amounts in these steps are for 30 × 106 cells, as the density separation will be carried out in a 15-ml tube and loaded with 30 × 106 cells. For higher numbers of cells, you will need to prepare multiple tubes in an identical manner. For ease of visualization of the different layers, make the alternating densities of Percoll with 10× HBSS with (Cellgro, cat. no. 20-021-cv) and without (Invitrogen, cat. no. 14065-056) phenol red. Initially, two solutions of 100% Percoll are produced, one clear and one red. Prepare the density gradients as directed in Table 1E.5.1 by using different-colored 100% Percoll in the adjacent layers, as shown in Figure 1E.5.7.

7. Prepare 30%, 42%, 52%, and 70% (v/v) Percoll concentrations by diluting the 100% Percoll with HBSS (1×), according to Table 1E.5.1.

Carry out density separation 8. In a 15-ml conical tube, gently layer Percoll gradients as follows: 1 ml 70% Percoll, 3 ml 52%, 4 ml 42%, and 5 ml 30% Percoll, from the bottom upward (Fig. 1E.5.7A). 9. Gently load 1 ml of the cell suspension from step 4 on top of the gradient (Fig. 1E.5.7A). 10. Centrifuge 30 min at ∼1000 × g, 4◦ C. Stop without using the brake. hAMSC cells will be in the top fraction (1), while hAECs cells will stay in the bottom one (4). The two intermediate fractions (2 and 3) will contain a mixture of the two cell types (Figs. 1E.5.7B and 1E.5.8).

11. If hAECs are not needed, carefully collect the hAMSCs in fraction 1 and transfer them into a new 15-ml tube. Discard the other fractions. 12. Fill up the tube with 1× HBSS, mix, and centrifuge for 5 min at ∼200 × g, 4◦ C. 13. Discard the supernatant and resuspend the pellet in 10 ml standard culture medium. 14. Count the cell number (UNIT 1C.3). 15. If cells are to be cultured, plate up to 1 × 105 cells per cm2 in standard culture medium. See Basic Protocol, step 21 for culture conditions. Table 1E.5.1 Preparation of Percoll Solutions for Isolation of hAMSCs

Percoll concentration

Isolation of Amniotic Mesenchymal Stem Cells

Total vol. 100% Percoll (ml) (ml)

HBSS (ml)

30%

5

1.5

3.5

42%

4

1.68

2.32

52%

3

1.52

1.48

70%

1

0.7

0.3

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A

B

cell suspension

30%

Percoll gradient

1 hAMSC fraction

centrifuge 42% 2 hAMSC/hAEC fraction 52% 3 hAEC/hAMSC fraction 70% 4 hAEC fraction

Figure 1E.5.7 Schematic representation of density separation with Percoll gradients before (A) and after (B) centrifugation.

A

B

C

D

Figure 1E.5.8 Morphology of hAMSCs and hAECs cultured for 24 hr after Percoll density separation. (A) Fraction 1: hAMSC monolayer; (B) fraction 2: islands of hAECs (red arrows) on a hAMSC monolayer; (C) fraction 3: islands of hAMSCs (yellow arrows) on a hAEC monolayer; (D) fraction 4: hAEC monolayer.

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Digestion solution 500 ml EMEM (with 25 mM HEPES buffer without L-glutamine, with Earle’s BSS; Lonza, cat. no. 12-136F) supplemented with: 1 mg/ml Collagenase type IV (Sigma, cat. no. C5138) 25 μg/ml DNase I (Sigma, cat. no. DN25) To be prepared fresh in a volume proportional to the amount of tissue.

Standard culture medium 430 ml DMEM (Lonza, cat. no. 12-604F, with 4.5 g of glucose/liter) 5 ml sodium pyruvate (Invitrogen, cat. no. 11360-070) 50 ml 10% (v/v) heat-inactivated fetal bovine serum (Invitrogen, cat. no. 16141) 5 ml 100 mM non-essential amino acid (Invitrogen, cat. no. 11140-050) 5 ml 200 mM L-glutamine (Cellgro, cat. no. 25-005-CI) 500 μl 55 mM 2-mercaptoethanol (Invitrogen, cat. no. 21985-023) 5 ml antibiotics solution (Cellgro, cat. no. 30-004-CI) 10 ng/ml epidermal growth factor (EGF; Sigma, cat. no. E9644, human recombinant) Store up to 2 weeks at 4◦ C COMMENTARY Background Information

Isolation of Amniotic Mesenchymal Stem Cells

Amniotic membrane is a tissue of fetal origin, which develops from the epiblast by 8 days after fertilization and prior to gastrulation. This raises the possibility that cells isolated from this tissue might retain the plasticity of pregastrulation embryo cells. hAMSCs have a mesenchymal phenotypic profile comparable to that of bone marrow MSCs, and they have the ability to differentiate toward mesodermal lineages (Parolini et al., 2008). As assessed by flow cytometry analysis, freshly isolated hAMSCs express some of the markers commonly used for the detection of MSCs, such as CD44, CD73, and CD90 (Fig. 1E.5.9). Moreover, it has been recently reported that a multipotent side population with multilineage differentiation potential is present in the mesenchymal region of the amniotic membrane (Kobayashi et al., 2008). hAMSCs have been shown to express major cartilage components after chondrogenic induction, with deposition of collagen II after in vivo implantation into the abdominal muscle of mice (Wei et al., 2009). Since there is no maternal contribution to the amnion membrane, unlike MSCs isolated from chorion, amnion-derived MSCs are entirely fetal.

Critical Parameters and Troubleshooting There is some variability in the cell yield between individuals. The authors ascribe these differences to the quality of tissue that is received, but this has not been specifically tested. It is clear that the time between the delivery of the placenta and the start of the cell isolation is critical. The tissue should be maintained in a refrigerator as soon as possible. This time in the refrigerator is called the cold ischemic time. It is best to minimize the cold ischemic time to <3 hr, if possible. Successful cell isolations have been conducted even 24 hr following delivery of the placenta, but the cell yield suffers considerably, and may be reduced to 1/3 of normal. The initial viability is not always indicative of the long-term quality of the cells. In many cases, the initial viability is >75%, and yet the cells do not proliferate robustly in culture. In order to evaluate the homogeneity of the cell population after isolation, a characterization by flow cytometry can be useful to determine the quality of the cell preparation. We suggest to evaluate the expression of some of the major markers commonly used for the detection of MSCs, such as CD44, CD73 (SH3/SH4), or CD90 (Thy-1). It is also useful to exclude

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0 100

M1

72.2%

M1

0 101

102

103

104

CD44-PE

Events

59.7%

1024

Events

256

Events

256

88.8%

M1

0 100

101

102

103

CD90-APC

104

100

101

102

103

104

CD73-PE

Figure 1E.5.9 FACS analysis of MSC marker expression on hAMSCs immediately after isolation. The percentage of the positive fraction is relative to isotype controls.

the possibility of leukocyte contamination by detection of CD45, a common leukocyte antigen.

Miki, T., Lehmann, T., Cai, H., Stotz, D.B., and Strom, S.C. 2005. Stem cell characteristics of amniotic epithelial cells. Stem Cells 23:15491559.

Anticipated Results

Parolini, O., Alviano, F., Bagnara, G.P., Bilic, G., B¨uhring, H.J., Evangelista, M., Hennerbichier, S., Liu, B., Magatti, M., Mao, N., Miki, T., Marongiu, F., Nakajima, H., Nikaido, T., Portmann-Lanz, C.B., Sankar, V., Soncini, M., Stadler, G., Surbek, D., Takahashi, T.A., Redi, H., Sakuragawa, N., Wolbank, S., Zeisberger, S., Zisch, A., and Strom, S.C. 2008. Concise review: Isolation and characterization of cells from human term placenta: Outcome of the first International Workshop on Placenta Derived Stem Cells. Stem Cells 26:300311.

The yield of hAMSCs from one term placenta is 20–100 × 106 cells that will continue to proliferate to passage 15.

Time Considerations The total time from stripping of the tissue to the final cell pellet is ∼3 to 3.5 hr.

Literature Cited Kobayashi, M., Yakuwa, T., Sasaki, K., Sato, K., Kikuchi, K., Kamo, I., Yokoyama, Y., and Sakuragawa, N. 2008. Multilineage potential of side population cells from human amnion mesenchymal layer. Cell Transplant. 17:291-301. Magatti, M., De Murani, S., Vertua, E., Gibelli, L., Wengler, G.S., and Parolini, O. 2008. Human amnion mesenchyme harbors cells with allogeneic T-cell suppression and stimulation capabilities. Stem Cells 26:182-192. Miki, T. and Strom, S.C. 2006. Amnion-derived pluripotent/multipotent stem cells. Stem Cell Rev. 2:133-142.

Soncini, M., Vertua, E., Gibelli, L., Zorzi, F., Denegri, M., Albertini, A., Wengler, G.S., and Parolini, O. 2007. Isolation and characterization of mesenchymal cells from human fetal membranes. J. Tissue Eng. Regen. Med. 1:296305. Wei, J.P., Nawata, M., Wakitani, S., Kametani, K., Ota, M., Toda, A., Konishi, I., Ebara, S., and Nikaido, T. 2009. Human amniotic mesenchymal cells differentiate into chondrocytes. Cloning Stem Cells 11:19-26.

Embryonic and Extraembryonic Stem Cells

1E.5.11 Current Protocols in Stem Cell Biology

Supplement 12

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

UNIT 1E.6

Sean Murphy,1,2 Sharina Rosli,1 Rutu Acharya,1 Louisa Mathias,2 Rebecca Lim,1 Euan Wallace,1 and Graham Jenkin2 1

Department of Obstetrics and Gynaecology, Monash University, Clayton, Victoria, Australia 2 Monash Immunology and Stem Cell Laboratories, Monash University, Clayton, Victoria, Australia

ABSTRACT Human amnion epithelial cells (hAECs) are a heterologous population positive for stem cell markers; they display multilineage differentiation potential, differentiating into cells of the endoderm (liver, lung epithelium), mesoderm (bone, fat), and ectoderm (neural cells). They have a low immunogenic profile and possess potent immunosuppressive properties. Hence, hAECs may be a valuable source of cells for cell therapy. This unit describes an efficient and effective method of hAEC isolation, culture, and cryopreservation that is animal product–free and in accordance with current guidelines on preparation of cells for clinical use. Cells isolated using this method were characterized after 5 passages by analysis of karyotype, cell cycle distribution, and changes in telomere length. The differentiation potential of hAECs isolated using this animal product–free method was demonstrated by differentiation into lineages of the three primary germ layers and expression of lineage-specific markers analyzed by PCR, immunocytochemistry, and C 2010 by John Wiley & histology. Curr. Protoc. Stem Cell Biol. 13:1E.6.1-1E.6.25.  Sons, Inc. Keywords: isolation r amniotic r amnion r epithelial r stem cells r cryopreservation r culture r characterization r placenta

INTRODUCTION Regenerative medicine is a relatively new field of medicine with the objectives of either aiding normal healing processes to be more efficient, or using special materials to regrow lost or damaged tissue. In addition to the current therapeutic strategies, there is an emerging interest in regenerative medicine as a potential alternative to complicated tissue/organ transplantation. Some of these methods are based on the use of stem cells to generate biological substitutes for damaged tissue/organs, thus avoiding some of the difficulties associated with transplantation, such as the lack of available donor tissue or the risks of recipient immune rejection of the donor transplant. Thus, the isolation and successful establishment of human embryonic stem cell (hESC) lines have been an important and most promising development in the field of regenerative medicine (Thomson et al., 1998; Reubinoff et al., 2000). However, despite their therapeutic potential, considerable legal and ethical challenges surround the derivation of these stem cell lines, necessitating consideration of alternative sources of stem cells for use in regenerative therapies. Recently, human amnion epithelial cells (hAECs) have attracted attention as a potential cell source for regenerative therapies (Parolini et al., 2008), with reports that these epithelial cells derived from human term amnion possess multipotent differentiation ability (Ilancheran et al., 2007), low immunogenicity (Bailo et al., 2004), and anti-inflammatory functions (Li et al., 2005). Importantly, while hAECs have similar Current Protocols in Stem Cell Biology 1E.6.1-1E.6.25 Published online April 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01e06s13 C 2010 John Wiley & Sons, Inc. Copyright 

Embryonic and Extraembryonic Stem Cells

1E.6.1 Supplement 13

properties to embryonic stem cells, they are not sourced from human embryos, as the amnion is usually discarded as medical waste along with other placental tissues following birth. In theory, hAECs could be isolated from every placenta, with almost 300,000 births per year in Australia (Laws and Hilder, 2008). Thus, once technology is established to isolate and store hAECs cost-effectively, it should be possible to create amnion cell banks encompassing all major histocompatability complex (MHC) immunotypes, which could be used for allogenic clinical applications. This potential led us to develop protocols for hAECs isolation that could be used in future in vivo regenerative therapies. We aimed to develop an efficient and effective method of hAEC isolation, culture, and cryopreservation that was animal product–free and in accordance with current guidelines on preparation of cells for clinical use (Therapeutic Goods Administration, 1989). Basic Protocol 1 describes an animal product–free cell isolation method we developed and compared to a previously published research method (UNIT 1E.3) that uses reagents containing animal products. We compared total cell yield, cell viability (trypan blue exclusion), and isolate cell purity (FACS analysis for EpCam+ cells). Basic Protocol 2 describes a method of animal product–free cell cyropreservation and thawing developed to support the long-term storage of hAECs that would be required for cell banking. Two animal product–free cryopreservation media, Cryostor CS5 and Synth-a-Freeze, were compared to a widely used serum-based medium. Post-thaw viability was measured by trypan blue exclusion and the recovery of cell metabolism determined using the CellTiter 96 AQueous One Solution Cell Proliferation Assay. Animal product–free culture conditions were developed using commercially available media (EpiLife, DMEM/F12) and human recombinant growth factors. We characterized hAECs when freshly isolated and after culture in animal product–free culture conditions. Freshly isolated hAECs were interrogated with a panel of monoclonal antibodies and analyzed by flow cytometry (Support Protocol 1). This analysis was then repeated on hAECs from the same amnion following culture for 5 passages. Karyotype, cell cycle, and telomere length stability of hAECs were analyzed after 5 passages by Giemsa band karyogram, cell cycle distribution, and by telomere length assay. Lastly, the differentiation potential of hAECs isolated using our animal product–free method was demonstrated by culturing hAECs in media developed to differentiate and maintain neuronal, osteogenic, adipogenic, and small airway epithelial cells (Support Protocol 2). Expression of markers associated with these lineages by these cells was determined by PCR. Assays for proliferation (Support Protocols 3,4) and telomere length (Support Protocol 5) are included for characterization of hAECs. BASIC PROTOCOL 1

ISOLATION OF AMNION EPITHELIAL CELLS This protocol was adapted from a previously published protocol that used animal-derived reagents (UNIT 1E.3). Specifically, reagents are altered to optimize cell yields and viability and to replace animal-derived products with animal product–free reagents. Cells obtained using this method would comply with standards derived for use in human clinical trials under current Therapeutic Goods Administration requirements (1989). Expected yield can range from 80–160 × 106 cells with ∼80% viability (Fig. 1E.6.1).

Materials

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

Placentas Hanks’ Balanced Salt Solution (HBSS; Invitrogen, cat. no. 14175) TrypZean (animal product–free recombinant trypsin; Sigma-Aldrich, cat. no. T3449) Soybean trypsin inhibitor (see recipe)

1E.6.2 Supplement 13

Current Protocols in Stem Cell Biology

EpiLife growth medium (animal product–free medium; see recipe) Anti-EpCAM-PE antibody (BD Biosciences, cat. no. 347198) Monoclonal mouse anti-CD90-PeCy5 (BD Biosciences, cat. no. 555597) Monoclonal mouse anti-CD105-APC (eBioscience, cat. no. 171057) 100- to 200-ml specimen containers, sterile Sterile scissors and forceps 15-cm petri dishes 37◦ C shaking water bath 40- and 70-μm filters 15-ml centrifuge tubes BD FACS Canto flow cytometer Additional reagents and equipment for performing a cell count (UNIT 1C.3) NOTE: Placentas were collected from healthy women with singleton pregnancies undergoing elective caesarean section delivery at term. Women gave written, informed consent for the collection of their placenta. The collection, and subsequent use, of placentas was performed with approval from the Southern Health Human Research Ethics Committee.

100 50 0

100 80 60 40 20 0

Cell number

106)

10

Clinical method 85%

Live cells (

200 105 150

6

Live cells (%)

Live cells (

106)

Traditional method

200 120 150

106

Live cells (%)

A

100 50 0

Viability

100 80 60 40 20 0

Cell number

83%

Viability

B isotype Counts

Counts

isotype 94%

92%

EpCAM

EpCAM

C

1% CD105

CD105

1% R8

CD90

R8

CD90

Figure 1E.6.1 Comparison of the previously established animal product–containing method (UNIT 1E.3) and our method suitable for clinical use. (A) Comparison of average hAEC number and viability [mean stated above histograms, error bars denote SEM, n = 92 (standard), n = 19 (clinical)]. (B) Comparison of epithelial marker expression (EpCAM) (percentage of EpCAM+ cells stated). (C) Comparison of mesenchymal marker (CD90/CD105) expression (number of CD90+ CD105+ cells <1%).

Embryonic and Extraembryonic Stem Cells

1E.6.3 Current Protocols in Stem Cell Biology

Supplement 13

Collect amnion 1. Carefully wash blood from placenta and membranes using 200 to 300 ml sterile HBSS. Placentas are transported in sterile standard collection containers.

2. Manually strip the amnion membrane from the chorion membrane of the placenta immediately after delivery, starting from the outer edge of the membrane and working towards the umbilical cord. Be careful to remove any pieces of contaminating chorion membrane from the amnion.

3. Cut the amnion membrane ∼2 cm from the umbilical cord and place in 100 ml room temperature HBSS in the collection container.

Wash the amnion 4. Remove the amnion membrane from the collection container and transfer to a 15-cm petri dish. 5. Cut each amnion membrane into 3 to 4 pieces, discarding bloody or torn pieces. 6. Wash repeatedly in HBSS until amnion membrane is completely clear of blood. Any blood present may reduce the efficiency of the trypsin enzyme.

Digest the amnion 7. Transfer all pieces of amnion membrane into a sterile specimen container containing 50 ml of TrypZean solution. 8. Incubate for 15 min in a water bath at 37◦ C with gentle shaking to remove contaminating blood cells. 9. Remove the TrypZean solution and discard. 10. Add 100 ml TrypZean solution and incubate for 60 min in a water bath 37o C with gentle shaking (first digestion). 11. Remove the amnion membrane pieces and place into an empty sterile specimen container taking care to allow TrypZean solution to drain from each piece of membrane. 12. Pass the remaining cell solution through a 70-μm filter into a 50-ml centrifuge tube and centrifuge 10 min at 1000 × g, room temperature. 13. Resuspend the cell pellet in 4 ml HBSS containing 1 mg/ml soybean trypsin inhibitor. Gently pipet the mixture repeatedly in order to obtain a single-cell suspension.

Repeat the digestion 14. Add 100 ml of TrypZean solution to the amnion membrane pieces (step 11) and incubate 60 min in a water bath at 37◦ C with gentle shaking (second digestion). 15. Remove the amnion membrane pieces and place into an empty sterile specimen container taking care to allow TrypZean solution to drain from each piece of membrane. 16. Pass the remaining cell solution through a 70-μm filter into a 50-ml centrifuge tube and centrifuge 10 min at 1000 × g, room temperature. 17. Resuspend the cell pellet in 4 ml HBSS containing 1 mg/ml soybean trypsin inhibitor. Gently pipet the mixture repeatedly in order to obtain a single-cell suspension. Amnion Epithelial Cell Isolation and Characterization for Clinical Use

18. Pool the cells from the two digests and pass through a 40-μm filter.

1E.6.4 Supplement 13

Current Protocols in Stem Cell Biology

19. Add EpiLife growth medium to a final volume of 10 to 20 ml and perform cell counts (UNIT 1C.3).

Assess the purity of the cells The purity of the cell isolate, as defined by the percentage (%) of cells that are confirmed to be epithelial is expected to be 90% to 95%. Analysis is performed by fluorescentactivated cell sorting (FACS) using the following steps. Although it is preferable to use freshly isolated and cryopreserved cells for clinical applications, if desired, the sorting/purification of EpCAM positive cells can be performed using this method. 20. Make up appropriate antibody cocktails using anti-EpCAM-PE (1:2 dilution), antiCD90-PeCy5 (1:250), and anti-CD105-APC (1:100) in HBSS. 21. For analysis and setting of appropriate gates for FACS, stain 1 × 106 cells with 40 μl antibody cocktails containing isotype controls in sterile 1-ml microcentrifuge tubes (e.g., IgG1 isotype control-PE + anti-CD90-PeCy5 + anti-CD105-APC). Keep ∼1 × 106 cells unstained to allow for FACS set up. 22. For cell sorting, resuspend remaining cells in complete antibody cocktail at a concentration of 25 × 106 cells/ml. 23. Incubate cells in antibody cocktails 60 min at 4◦ C, protected from light. 24. Wash the cells three times, each time in 1 ml HBSS, centrifuge 5 min at 700 × g, room temperature, and resuspend at 5–10 × 106 cells/ml for FACS.

CRYOPRESERVATION AND THAWING OF hAECS Animal product–free cryopreservation media, Cryostor CS5 and Synth-a-Freeze, are comparable to a serum-based cryopreservation medium (FBS +10% DMSO), in their ability to support cryopreservation, long-term storage, and recovery of viable hAECs. Cryopreservation of hAECs is performed on both freshly isolated cells and after FACS, to store cells for future clinical applications.

BASIC PROTOCOL 2

Materials Freshly isolated or FACS-sorted hAECs (Basic Protocol 1) Cryopreservation medium (see recipe) Liquid nitrogen storage system EpiLife growth medium (animal product–free medium; see recipe) O-ring cryopreservation vials Mr. Frosty freezing container (Thermo Fisher Scientific, cat. no. C1562) −80◦ C freezer 37◦ C water bath 15-ml centrifuge tubes Additional reagents and equipment for counting the cells using trypan blue exclusion (UNIT 1C.3) Cryopreserve the cells 1. Count the freshly isolated or FACS-sorted cells and determine viability by trypan blue exclusion (UNIT 1C.3). 2. Centrifuge 5 min at 700 × g, room temperature. 3. Resuspend the cells at 5 × 106 cells/ml in cryopreservation medium. Embryonic and Extraembryonic Stem Cells

1E.6.5 Current Protocols in Stem Cell Biology

Supplement 13

4. Pipet 1 ml cells per O-ringed cryopreservation vial and put the vials into precooled a Mr. Frosty freezing container. Mr. Frosty freezing containers should be cooled to 4◦ C several hours prior to use.

5. Place the container for 24 to 48 hr at −80◦ C. 6. Transfer the vials into liquid nitrogen. The vials can be stored indefinitely in liquid nitrogen.

Thaw and assess viability of the cells 7. Remove the cryopreservation vials from the liquid nitrogen and place on ice. 8. Thaw the cells in water bath at 37◦ C until only a small piece (5 × 5–mm) of ice remains. It is important that the cell solution does not warm to reduce the toxic effect of DMSO in the cryopreservation medium.

9. Transfer the cells into a 15-ml centrifuge tube and add cold (4◦ C) EpiLife growth medium dropwise to a final volume of 15 ml. 10. Centrifuge 5 min at 700 × g in a centrifuge cooled to 4◦ C. 11. Resuspend the cells in 5 ml EpiLife growth medium, perform cell counts, and analyze viability by trypan blue exclusion (UNIT 1C.3). Expected viability post-thaw will be 5% to 10% less than pre-cryopreservation viability.

12. Cells can be used at this stage for clinical or experimental purposes. Alternatively, cells can be plated at 2.6 × 104 cells/cm2 in EpiLife growth medium supplemented with human recombinant growth factors. 13. Maintain hAECs at 37◦ C, 5% CO2 , changing the medium every 3 days until confluent for 5 passages. These cells are then characterized by Giemsa band karyogram analysis, cell cycle distribution, and by measuring changes to telomere length (Support Protocol 5). The average telomere length of freshly isolated cells is compared to cells of the same isolate after 5 passages.

SUPPORT PROTOCOL 1

CHARACTERIZATION OF hAECS BY FLOW CYTOMETRY Freshly isolated hAECs are stained with a panel of monoclonal antibodies and analyzed by flow cytometry. A further group of cells are cultured for 5 passages and then stained with the same panel of antibodies and analyzed by flow cytometry.

Materials Fresh isolates of hAECs (Basic Protocol 1) or cultures of hAECs after 5 passages (Basic Protocol 2) FACS buffer (see recipe) Panel of monoclonal antibodies (Table 1E.6.1) provided by BD Biosciences Amnion Epithelial Cell Isolation and Characterization for Clinical Use

96-well tissue culture plates BD FACS Calibur flow cytometer

1E.6.6 Supplement 13

Current Protocols in Stem Cell Biology

1. Resuspend the cells at 1 × 106 cells/ml in FACS buffer. 2. Add 50 μl of cell suspension (50,000 cells) into each well of a 96-well plate. 3. Pipet 1 μl of monoclonal antibody or isotype control to each well (primary antibody). 4. Incubate 30 min at 4◦ C. 5. Add 200 μl of FACS buffer and centrifuge 3 min at 400 × g, room temperature. Remove the supernatant carefully without dislodging the pellet. 6. Add 30 μl secondary antibody and incubate 30 min at room temperature. 7. Add 200 μl of FACS buffer and centrifuge 3 min at 400 × g, room temperature. Remove the supernatant carefully without dislodging pellet. 8. Add 50 μl of FACS buffer and analyze with BD FACS Calibur flow cytometer. An example of the expected FACS plot for HLA-DQ is shown in Figure 1E.6.2.

250

Gated events: 27225

SSC-Height

R1

Region

% Total

R1

96.24

0 0

50

100 150 FSC-Height

200

250

104

Gate: G1

3

hAEC P0 HLA-DQ

FL4APC

10

Quad

% Gated

102

UL

0.02

101

UR

0.11

LL

84.06

LR

15.81

100

100

101

102

103

104

FL1 FITC

104

Gate: G1

3

hAEC P5 HLA-DQ

FL4APC

10

Quad

% Gated

102

UL

0.04

101

UR

0.19

LL

28.77

100

LR

71.01

100

101

102

103

104

FL1 FITC

Figure 1E.6.2 FACS plot of hAECs stained for HLA-DQ. Freshly isolated hAECs were stained with HLA-DQ and analyzed by flow cytometry. Cells from the same amnion were cultured for 5 passages, stained with HLA-DQ, and analyzed by flow cytometry.

Embryonic and Extraembryonic Stem Cells

1E.6.7 Current Protocols in Stem Cell Biology

Supplement 13

Table 1E.6.1 List of Monoclonal Antibodies Used to Interrogate hAECs

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

BD mAb

Host

Isotype

CD1a

Mouse

IgG1

CD1b

Mouse

IgG1

CD1d

Mouse

IgG1

CD2

Mouse

IgG1

CD3

Mouse

IgG2a

CD4

Mouse

IgG1

CD5

Mouse

IgG1

CD6

Mouse

IgG1

CD7

Mouse

IgG1

CD8

Mouse

IgG1

CD9

Mouse

IgG1

CD10

Mouse

IgG1

CD11a

Mouse

IgG1

CD11b

Mouse

IgG1

CD11c

Mouse

IgG1

CD13

Mouse

IgG1

CD14

Mouse

IgG1

CD15

Mouse

IgM

CD15s

Mouse

IgG1

CD16

Mouse

IgG1

CD18

Mouse

IgG1

CD19

Mouse

IgG1

CD20

Mouse

IgG2b

CD21

Mouse

IgG1

CD22

Mouse

IgG1

CD23

Mouse

IgG1

CD24

Mouse

IgG2a

CD25

Mouse

IgG1

CD26

Mouse

IgG1

CD27

Mouse

IgG1

CD28

Mouse

IgG1

CD29

Mouse

IgG2a

CD30

Mouse

IgG1

CD31

Mouse

IgG1

CD32

Mouse

IgG2b

CD33

Mouse

IgG1

CD34

Mouse

IgG1

CD35

Mouse

IgG1

CD36

Mouse

IgM continued

1E.6.8 Supplement 13

Current Protocols in Stem Cell Biology

Table 1E.6.1 List of Monoclonal Antibodies Used to Interrogate hAECs, continued

BD mAb

Host

Isotype

CD37

Mouse

IgG1

CD38

Mouse

IgG1

CD40

Mouse

IgG1

CD41a

Mouse

IgG1

CD41b

Mouse

IgG3

CD42a

Mouse

IgG1

CD42b

Mouse

IgG1

CD43

Mouse

IgG1

CD44

Mouse

IgG2b

CD45

Mouse

IgG1

CD45RA

Mouse

IgG2b

CD45RB

Mouse

IgG1

CD45RO

Mouse

IgG2a

CD46

Mouse

IgG2a

CD47

Mouse

IgG1

CD48

Mouse

IgM

CD49a

Mouse

IgG1

CD49b (1)

Mouse

IgG1

CD49b (2)

Mouse

IgG2a

CD49c

Mouse

IgG1

CD49d

Mouse

IgG1

CD49e (1)

Mouse

IgG1

CD49e (2)

Mouse

IgG2a

CD49f

Mouse

IgG2b

CD50

Mouse

IgG1

CD51/61

Mouse

IgG1

CD53

Mouse

IgG1

CD54

Mouse

IgG1

CD55

Mouse

IgG2a

CD56

Mouse

IgG2b

CD57

Mouse

IgM

CD58

Mouse

IgG2a

CD59

Mouse

IgG2a

CD61

Mouse

IgG1

CD62e

Mouse

IgG1

CD62L

Mouse

IgG1

CD62P

Mouse

IgG1

CD63

Mouse

IgG1

CD64

Mouse

IgG1 continued

Embryonic and Extraembryonic Stem Cells

1E.6.9 Current Protocols in Stem Cell Biology

Supplement 13

Table 1E.6.1 List of Monoclonal Antibodies Used to Interrogate hAECs, continued

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

BD mAb

Host

Isotype

CD66

Mouse

IgG1

CD66b

Mouse

IgM

CD66f

Mouse

IgG1

CD69

Mouse

IgG1

CD70

Mouse

IgG3

CD71

Mouse

IgG2a

CD72

Mouse

IgG2b

CD73

Mouse

IgG1

CD74

Mouse

IgG2a

CD77

Mouse

IgM

CD79b

Mouse

IgG1

CD80

Mouse

IgG1

CD81

Mouse

IgG1

CD83

Mouse

IgG1

CD84

Mouse

IgG1

CD85J

Mouse

IgG2b

CD86

Mouse

IgG1

CD87

Mouse

IgG1

CD88

Mouse

IgG1

CD89

Mouse

IgG1

CD90

Mouse

IgG1

CD91

Mouse

IgG1

CD94

Mouse

IgG1

CD95

Mouse

IgG1

CD97

Mouse

IgG1

CD98

Mouse

IgG1

CD99

Mouse

IgG2a

CD99R

Mouse

IgM

CD100

Mouse

IgG1

CD103

Mouse

IgG1

CD104

Mouse

IgG1

CD106

Mouse

IgG1

CD108

Mouse

IgG2a

CD109

Mouse

IgG1

CD110

Mouse

IgG1

CD117

Mouse

IgG1

CD123

Mouse

IgG1

CD134

Mouse

IgG1

CD135

Mouse

IgG1 continued

1E.6.10 Supplement 13

Current Protocols in Stem Cell Biology

Table 1E.6.1 List of Monoclonal Antibodies Used to Interrogate hAECs, continued

BD mAb

Host

Isotype

CD137 (1)

Mouse

IgG1

CD137 (2)

Mouse

IgG1

CD138

Mouse

IgG1

CD140a

Mouse

IgG2a

CD140b

Mouse

IgG2a

CD141

Mouse

IgG1

CD142

Mouse

IgG1

CD146

Mouse

IgG1

CD147

Mouse

IgG1

CD150

Mouse

IgG1

CD151

Mouse

IgG1

CD152

Mouse

IgG2a

CD153

Mouse

IgG1

CD154

Mouse

IgG1

CD158a

Mouse

IgM

CD158b

Mouse

IgG2b

CD161

Mouse

IgG1

CD162

Mouse

IgG1

CD163

Mouse

IgG1

CD164

Mouse

IgG2a

CD165

Mouse

IgG1

CD166

Mouse

IgG1

CD172b

Mouse

IgG1

CD177

Mouse

IgG1

CD180

Mouse

IgG2a

CD181

Mouse

IgG2a

CD182

Mouse

IgG1

CD183

Mouse

IgG1

CD184

Mouse

IgG2b

CD195

Mouse

IgG1

CD200

Mouse

IgG1

CD201

Rat

IgG1

CD206

Mouse

IgG1

CD209

Mouse

IgG2b

CD210

Rat

IgG1

CD212

Rat

IgG1

CD220

Mouse

IgG1

CD221

Mouse

IgG1

CD226

Mouse

IgG1

CD227

Mouse

IgG1 continued

Current Protocols in Stem Cell Biology

Embryonic and Extraembryonic Stem Cells

1E.6.11 Supplement 13

Table 1E.6.1 List of Monoclonal Antibodies Used to Interrogate hAECs, continued

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

BD mAb

Host

Isotype

CD229

Mouse

IgG1

CD235a

Mouse

IgG2b

CD244

Mouse

IgG1

B7-H2

Mouse

IgG2b

CMRF-44

Mouse

IgG1

CMRF-5

Mouse

IgG1

B2-MICROGLOBULIN

Mouse

IgG1

CLIP

Mouse

IgG1

gd-TCR

Mouse

IgG1

Vb8-TCR

Mouse

IgG1

CLA

Rat

IgG1

EGF-R

Mouse

IgG2a

fMLP-R

Mouse

IgG1

fII-R

Mouse

IgG1

HLA-ABC

Mouse

IgG1

HLA-A2

Mouse

IgG2b

HLA-DQ

Mouse

IgG2a

HLA-DR

Mouse

IgG2a

HLADRDPD

Mouse

IgG2a

abTCR

Mouse

IgM

IntegrinB7

Mouse

IgG2b

M-calpain

Mouse

IgG1

LAIR-1

Mouse

IgG1

NL-B1

Mouse

IgG1

NK-G2d

Mouse

IgG1

NK-P46

Mouse

IgG1

ABC-G2

Mouse

IgG2b

Blood group A

Mouse

IgG3

NP<-ALK/AL

Mouse

IgG3

B glycoprotein

Mouse

IgG2b

Invariant NK

Mouse

IgG1

MUC2

Mouse

IgG1

NGF-R

Mouse

IgG1

PRR2

Mouse

IgG1

Siglec-6

Mouse

IgG1

Siglec-7

Mouse

IgG1

B5-TCR

Mouse

IgG2a

Common gamma chain

Rat

IgG2b

Stro-1

Mouse

IgG1

CD105

Mouse

IgG1

1E.6.12 Supplement 13

Current Protocols in Stem Cell Biology

DIFFERENTIATION OF hAECS Differentiation potential of hAECs isolated using Basic Protocol 1 was demonstrated by the maintenance of hAECs under conditions designed to promote differentiation along cell lineages representing the three primary germ layers. Neural, lung epithelial, adipogenic, and osteogenic differentiation were induced using the following protocol.

SUPPORT PROTOCOL 2

NOTE: As differentiation media may contain animal products, cells maintained in these media would no longer be suitable for clinical applications.

Materials Freshly isolated hAEC (Basic Protocol 1) EpiLife growth medium (see recipe) EpiLife coating matrix (Invitrogen, cat. no. R-011-K) Neural differentiation medium (see recipe) Small airway growth medium (see recipe) Adipogenic differentiation medium (see recipe) Osteogenic differentiation medium (see recipe) Hanks’ Balanced Salt Solution (HBSS; Invitrogen, cat. no. 14175) Poly-D-lysine/laminin-coated glass coverslips (BD Biosciences, cat. no. 354087) or 12-well multiwell plates (BD Biosciences, cat. no. 351143) 1. Following isolation of hAECs using Basic Protocol 1, sort EpCAM+ CD90− CD105− cells. 2a. For neural differentiation: Plate cells in EpiLife growth medium at a density of 10,000 cells/cm2 on poly-D-lysine/laminin-coated coverslips or other suitable growth surface to promote attachment of neural progenitor cells. 2b. For small airway epithelial cell differentiation: Plate cells in EpiLife growth medium at a density of 10,000 cells/cm2 on coverslips coated with EpiLife coating matrix. 2c. For adipogenic and osteogenic differentiation: Plate cells in EpiLife growth medium at a density of 20,000 cells/cm2 on coverslips coated with EpiLife coating matrix. 3. Allow cells to attach for 72 hr before medium change. 4a. For neural differentiation: Change medium to neural differentiation medium. 4b. For lung cell differentiation: Change medium to the small airway growth medium. 4c. For adipogenic differentiation: Allow cells to achieve 100% confluence, and then change medium to adipogenic differentiation medium. 4d. For osteogenic differentiation: Allow cells to achieve 100% confluence, and then change medium to osteogenic differentiation medium. 5. After an additional 72 hr, wash with 1 ml HBSS and replace medium every 3 days for 28 days. For control groups, continue replacing media every 2 to 3 days with EpiLife growth medium. Replicate wells can be used to perform assays to characterize hAEC differentiation. In this case, immunocytochemistry was performed to detect GFAD, MAP2, and SP-C—markers of glial cells, neurons, and lung epithelium, respectively. Oil Red-O and Alizarin Red-S histological stains were used to demonstrate presence of lipid and mineralized calcium, respectively. Polymerase chase reaction was used to demonstrate induction of lineage-specific gene expansion. Primer sequence and cycling conditions are stated in Table 1E.6.2.

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Table 1E.6.2 Primer Sequences and Conditions for Characterizing hAECs by PCR

Gene

Primer sequence

Nestin

Forward: CAGCTGGCGCACCTCAAGATG

Annealing temperature

Cycles

60◦ C

40

58◦ C

40

56◦ C

45

60◦ C

45

58◦ C

35

59◦ C

35

53◦ C

35

60◦ C

35

63◦ C

35

60◦ C

35

60◦ C

35

Reverse: AGGGAAGTTGGGCTCAGGACTGG β - 3 - Tubulin

Forward: CTCAGGGGCCTTTGGACATC Reverse: CAGGCAGTCGCAGTTTTCAC

SP-A

Forward: AAGCCACACTCCACGACTTTAGA Reverse: GCCCATTGCTGGAGAAGACCT

SP-B

Forward: CATCGACTACTTCCAGAACCAGAC Reverse: GCAGATTGCCGCCCGCCACCAGAGG

SP-C

Forward: TTGGTCCTTCACCTCTGTCC Reverse: CTCCAGAACCATCTCCGTGT

SP-D

Forward: TCATGTGTAGCTCAGTGGAGAGTG Reverse: AGGTTCTCCAACAGAGCCATTGT

Osteocalcin

Forward: GTGCAGAGTCCAGCAAAGGT Reverse: CTGGAGAGGAGCAGAACTGG

Osteonectin

Forward: GTGCAGAGGAAACCGAAGAG Reverse: AAGTGGCAGGAAGAGTCGAA

LPL

Forward: ATGGAGAGCAAAGCCCTGCTC Reverse: TACAGGGCGGCCACAAGTTTT

PPARγ

Forward: GCTGTTATGGGTGAAACTCTG Reverse: ATAAGGTGGAGATGCAGGCTC

GAPDH

Forward: CCTCAAAGGCATCCTGGGCTACAC Reverse: CATGTGGGCCATGAGGTCCACCAG

SUPPORT PROTOCOL 3

CellTiter 96 AQUEOUS ONE SOLUTION CELL PROLIFERATION ASSAY This assay measures the activity of enzymes that reduce MTS to formazan. These reductions take place only when reductase enzymes are active, and therefore conversion is often used as an indirect measure of cell proliferation. Changes in metabolic activity can give large changes in MTS results while the number of viable cells is constant. The CellTiter 96 AQueous One Solution Cell Proliferation Assay (the technical bulletin is #245 and can be found at http://www.promega.com/tbs/tb245/tb245.pdf) was used to determine the metabolic activity of hAECs following cryopreservation and thawing. This assay is performed following the manufacturer’s instructions. The steps are briefly described below.

Materials

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

Cryopreserved and thawed hAECs EpiLife growth medium (see recipe) CellTiter 96 AQueous One Solution Cell Proliferation Assay (Promega, cat. no. G3582) containing: Cell Titer 96 Aqueous One Solution Reagent

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96-well opaque-walled tissue culture plates compatible with fluorometer Multichannel pipettor Humidified 37◦ C, 5% CO2 incubator Fluorescence plate reader with 490-nm and 690-nm filters NOTE: Protect CellTiter 96 AQueous One Solution Reagent from direct light to prevent increased background readings. 1. Following isolation of hAECs (Basic Protocol 1), cells were cryopreserved in either FBS + 10% DMSO, Cryostor CS5, or Synth-a-Freeze. Thaw the cells according to Basic Protocol 2 and determine cell counts and viability by trypan blue excursion (UNIT 1C.3). 2. Set up 96-well assay plates in triplicate containing hAECs at 2–6 × 104 cells/cm2 in 100 μl of EpiLife growth medium. Set up triplicate wells without cells to serve as a negative control to determine background fluorescence. 3. Culture cells in a humidified 37◦ C, 5% CO2 incubator for 24 hr. 4. Remove assay plates from the incubator and add 20 μl/well CellTiter 96 AQueous One Solution Reagent. 5. Incubate using standard cell culture conditions for 1 to 4 hr. 6. Shake plates for 10 sec and record fluorescence at 490 nm (reference wavelength: 690 nm). The presence of serum in cryopreservation media does not improve hAEC metabolism 24 hr post-thaw. Metabolic recovery was greatest when hAECs were cryopreserved in Cryostor CS5 (Fig. 1E.6.3B).

A

B

Post-thaw viability

Post-thaw metabolism 2.0

*

**

1.5

80 OD490

60 40

1.0 0.5

20

e

r

fre ez

to os

ahnt Sy

ahnt Sy

ry

ez fre

os ry C

C

r to

SO D M

SO

0.0

e

0

D M

Percent recovery

100

* P 0.001 ** P 0.01

Figure 1E.6.3 Comparison of several serum-free cryopreservation and culture media. (A) Postthaw viability of hAECs cryopreserved in animal product–free cryopreservation media Cryostor CS5, Synth-a-Freeze, and a commonly used serum-based media (FBS + 10% DMSO), expressed as percentage live cell recovery (ANOVA, error bars denote SEM, n = 10). (B) MTS assay measuring hAEC metabolism 7 days post-thaw. Cryostor CS5 was shown to be the preferred cryopreservation media for cell recovery and maintenance in vitro (ANOVA, error bars denote SEM, n = 20).

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SUPPORT PROTOCOL 4

CELL PROLIFERATION ELISA To determine which culture medium best supports hAEC proliferation, a colorimetric Cell Proliferation ELISA, BrdU is performed, according to the manufacturer’s instructions. The following is a brief description of the assay.

Materials Cultures of hAEC Appropriate culture medium Cell Proliferation ELISA, BrdU (colorimetric; Roche, cat. no. 11647229001) containing: BrdU Labeling Reagent FixDenat (ready to use) Anti-BrdU-POD Antibody Dilution Solution (ready to use) Washing Buffer PBS, 10× Substrate Solution TMB (ready to use) 96-well opaque-walled tissue culture plates compatible with fluorometer Multichannel pipettor 37◦ C, 5% CO2 humidified incubator Fluorescence plate reader with 370-nm and 490-nm filters Additional reagents and equipment for cell counting (UNIT 1C.3) Prepare for the assay 1. Prepare working solutions as per the manufacturer’s instructions. 2. Set up 96-well assay plates in triplicate containing hAECs at 2–6 × 104 cells/cm2 in the appropriate culture medium. (Set up triplicate wells without cells to determine background fluorescence).

Start the assay 3. Culture cells in a humidified 37◦ C, 5% CO2 incubator for 3 to 14 days. Longer incubation periods can be used if desired.

Label the cells 4. Add 10 μl/well BrdU labeling reagent (final concentration 10 μM BrdU) and incubate cells for an additional 24 hr in a humidified 37◦ C, 5% CO2 incubator. 5. Remove the labeling solution by aspiration.

Fix the cells 6. Add 200 μl/well FixDenat solution (from the kit) and incubate 30 min at 15◦ to 25◦ C. 7. Remove the FixDenat solution by aspiration.

Detect labeling 8. Add 100 μl/well anti-BrdU-POD working solution (from the kit) and incubate 90 min at 15◦ to 25◦ C. 9. Remove the antibody conjugate by inverting the plate and flicking.

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

10. Rinse the wells three times, each time with 300 μl/well washing buffer (from the kit), inverting, and flicking between washes. 11. Remove the washing buffer by aspiration.

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12. Add 100 μl/well substrate solution and incubate at 15◦ to 25◦ C for 30 min or until color development is sufficient for photometric detection. 13. Measure the absorbance of the samples in an ELISA reader at 370 nm (reference wavelength: 490 nm). While hAECs cultured in the presence of serum may have a greater proliferation rate compared to serum-free medium, the optimization of growth factors in serum-free medium should result in similar proliferation ratios.

TELOMERASE ASSAY TO ASSESS TELOMERE LENGTH An assay to determine the telomere length of freshly isolated hAECs is performed using the TeloTAGGG Telomere Length Assay, according to the manufacturer’s instructions. A further group of cells were maintained under animal product–free culture conditions for 5 passages and analyzed using the same assay.

SUPPORT PROTOCOL 5

Materials hAEC cells, freshly isolated and/or cultured for 5 passages Kit or reagents for genomic DNA isolation (phenol, chloroform, sodium acetate, ethanol) TeloTAGGG Telomere Length Assay (Roche, cat. no. 12209136001) containing: HinfI RsaI Digestion buffer, 10× Water, nuclease free Control DNA DIG molecular weight marker Loading buffer DIG Easy Hyb granules Telomerase probe Washing buffer Maleic acid buffer Blocking buffer Anti-DIG AP Detection buffer Substrate solution (CDP-Star, ready to use) HCl solution (0.25 M HCl) Denaturation solution (0.5 M NaOH, 1.5 M NaCl) Neutralization solution (0.5 M Tris·Cl, pH 7.5, 3 M NaCl) Stringent wash buffer I (2× SSC, 0.1% SDS) Stringent wash buffer II (0.2× SSC, 0.1% SDS) 42◦ C shaking incubator Absorbent paper Hybridization bag Bio-Rad ChemiDoc Densitometer Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2000) and Southern blotting (Brown, 1999) Prepare genomic DNA 1. Isolate genomic DNA from 1 × 106 freshly isolated hAECs, as well as after passage 5. 2. Resuspend 1 μg of genomic DNA in a total volume of 17 μl nuclease-free H2 O.

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3. Prepare HinfI/RsaI enzyme mixture by mixing equal volumes of each enzyme to result in a final concentration of 20 U/μl for each enzyme.

Digest DNA 4. Digest genomic DNA by adding 2 μl of 10× digestion buffer and 1 μl of HinfI/RsaI enzyme mixture and incubating for 2 hr at 37◦ C. 5. Stop the reaction by adding 4 μl of 5× gel electrophoresis loading buffer.

Electrophorese digested DNA 6. Separate the digested genomic DNA by agarose gel electrophoresis (Voytas, 2000) on a 0.8% agarose gel, loading 1 μg of genomic DNA per well. 7. Load 10 μl of DIG molecular weight marker to either side of genomic DNA samples and run the gel at 5 V/cm in 1× TAE buffer.

Blot the gel 8. Submerge the agarose gel in 250 ml HCl solution for 5 to 10 min until the bromphenol blue stain changes its color to yellow. 9. Rinse the gel two times, each time in 250 ml H2 O. 10. Submerge the gel in 250 ml denaturation solution twice for 15 min each at room temperature. 11. Rinse the gel two times, each time in 250 ml H2 O. 12. Submerge the gel in 250 ml neutralization solution twice for 15 min at room temperature. 13. Perform a Southern blot (Brown, 1999) to transfer genomic DNA to a positively charged nylon membrane by capillary action, at room temperature using 20× SSC solution as a transfer buffer. For maximum sensitivity, perform the transfer overnight. 14. Fix the transferred DNA to the wet blotting membrane by baking the membrane at 120◦ C for 20 min.

Hybridize the DNA with telomerase probe 15. Wash the blotting membrane with 2× SSC solution. 16. Prewarm 25 ml DIG Easy Hyb solution to 42◦ C. 17. Submerge the blot in 18 ml DIG Easy Hyb solution and incubate for 30 to 60 min at 42◦ C with gentle agitation. 18. Prepare the hybridization solution by adding 1 μl telomere probe per 5 ml prewarmed DIG Easy Hyb solution and mix. 19. Discard the DIG Easy Hyb solution and add 6.5 ml hybridization solution to the membrane and incubate for 3 hr at 42◦ C with gentle agitation.

Wash the membrane 20. Wash the membrane two times, each time with 250 ml stringent washing buffer I for 5 min at room temperature. 21. Wash the membrane two times, each time with 250 ml stringent wash buffer II for 15 to 20 min at 50◦ C. Amnion Epithelial Cell Isolation and Characterization for Clinical Use

22. Rinse the membrane in at least 100 ml washing buffer (from the kit) for 1 to 5 min at room temperature.

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Visualize telomere length 23. Incubate the membrane in 100 ml blocking buffer for 30 min at room temperature. 24. Incubate the membrane in 50 to 100 ml Anti-DIG-AP working solution for 30 min at room temperature. 25. Wash the membrane two times for 15 min each with 200 ml washing buffer at room temperature. 26. Incubate the membrane in 100 ml detection buffer for 2 to 5 min at room temperature. 27. Discard the detection buffer and pour off excess liquid by placing the membrane DNA side up on a sheet of absorbent paper. Do not let the membrane dry. 28. Immediately place the wet membrane into a open hybridization bag and apply 3 ml of substrate solution homogeneously over the membrane. 29. Incubate the membrane for 5 min at room temperature. 30. Squeeze out excess substrate solution and seal the edges of the hybridization bag. 31. Expose using a Bio-Rad ChemiDoc for 5 to 20 min at room temperature. 32. Calculate the mean telomere length by analyzing the image with a densitometer. To obtain reliable results, the signal strength must be within the linear range of the imaging system.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Adipogenic differentiation medium 200 ml complete culture medium: a-MEM (Invitrogen, cat. no. A10490-01) + 16.5% (v/v) FBS 0.5 μM dexamethasone (100 μl of 1 mM stock in deionized water) 0.5 μM isobutylmethylxanthine (20 μl of 5 mM stock in methanol) 50 μM indomethacin (333 μl of 30 mM stock in methanol) Store up to 4 weeks at 4◦ C Cryopreservation medium Fetal bovine serum + 10% (v/v) DMSO Cryostor CS5 (Biolife Solutions, cat. no. 610201) Synth-a-Freeze (Invitrogen, cat. no. R-005-50) EpiLife growth medium (animal product–free) 500 ml EpiLife basal medium (Invitrogen, cat. no. M-EPI-500-CA) 1% (v/v) supplement S7 (Invitrogen, cat. no. S7 S-017-5) Store up to 4 weeks at 4◦ C FACS buffer 5 ml of 10% (w/v) bovine serum albumin [BSA; 0.1% (w/v) BSA] 1 ml of 10% (v/v) azide Make up to 500 ml with phosphate-buffered saline (PBS) Store up to 4 weeks at 4◦ C EDTA can be added to a concentration of 2 mM to prevent cell clumping.

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Neural differentiation medium 500 ml Neurobasal-A medium (Invitrogen, cat. no. 10888-022) 2% (v/v) B-27 Serum-Free supplement (Invitrogen, cat. no. 17504-044) Store up to 4 weeks at 4◦ C Oil Red-O, 0.5% (w/v), stock 2.5 g Oil Red O 500 ml isopropyl alcohol Dissolve completely Store in a tightly capped bottle up to 1 year at room temperature Oil Red-O working solution 3 parts 0.5% (w/v) Oil Red-O stock (see recipe) 2 parts Hanks’ Balanced Salt Solution (HBSS; Invitrogen, cat. no. 14175) Mix and wait 10 min Filter stain through sterile filter Wait 10 min before use Prepare fresh Osteogenic differentiation medium 192 ml complete culture medium: a-MEM (Invitrogen, cat. no. A10490-01) + 16.5% (v/v) FBS 10 nM dexamethasone (20 μl of a 1:10 dilution of 1 mM stock solution in deionized water) 20 mM β-glycerophosphate (8 ml of 0.5 M stock in complete culture medium) 50 μM L-ascorbic acid 2-phosphate (200 μl of 50 mM stock solution in deionized water) Store up to 4 weeks at 4◦ C Small airway growth medium 500 ml small airway basal medium (Lonza, cat. no. CC-3119) SAGM SingleQuot Kit supplement and growth factors (Lonza, cat. no. CC-4124) Store up to 4 weeks at 4◦ C Soybean trypsin inhibitor 8 mg soybean trypsin inhibitor (Sigma-Aldrich, cat. no. T6522) 8 ml Hanks’ Balanced Salt Solution (HBSS; Invitrogen, cat. no. 14175) Dissolve completely and filter through 0.22-μm filter before use Prepare fresh Stain for osteogenesis 1 g Alzarin Red S 100 ml deionized water Adjust pH of solution to 4.1 to 4.3 using 0.1% ammonium hydroxide Filter stain through sterile filter Store up to 3 months at room temperature COMMENTARY Background Information Amnion Epithelial Cell Isolation and Characterization for Clinical Use

Following fertilization, the zygote (diploid cell derived from ovum from a female and a sperm cell from a male) undergoes a series

of cell divisions to form a solid ball of cells (the morula), and then a spherical layer of cells that surrounds a central fluid-filled cavity called the blastula. The blastocyst arises after

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compaction of the blastula and comprises an inner cell mass, which subsequently forms the embryo, and an outer cell mass, or trophoblast, which forms the placenta. In humans, 4 to 5 days post-fertilization the inner cell mass becomes differentiated into two tissues: the hypoblast (primary endoderm), which will form most extra-embryonic structures, and the epiblast (primary ectoderm) from which the embryo and some extra-embryonic structures will develop. Prior to gastrulation, epiblast cells migrate along the walls of the amniotic cavity and form the amnion epithelium. The migration of cells from the epiblast to the lining of the amniotic cavity occurs before the formation of the three primary germ layers. Due to their origin, it is believed that amnion epithelial cells may possess multipotent differentiation potential similar to that of ESC, which are derived from the inner cell mass. The potential of the amniotic membrane, per se, to treat pathologies has been recognized for some time. As early as 1910, the amnion membrane has been used to promote the healing of skin burns, and from the 1990s for ocular surface reconstruction (Trelford and Trelford-Sauder, 1979; Fernandes et al., 2005). Recently, cells derived from the human amnion membrane have been used in the treatment of various animal models of disease, such as for Parkinson’s Disease (Kakishita et al., 2000, 2003), spinal cord injury (Sankar and Muthusamy, 2003), and diabetes (Wei et al., 2003; Hou et al., 2008). The use of cells isolated from human term placenta for regenerative medicine represents a field of investigation that is still in its infancy, but holds great promises on several fronts. Specifically, their plasticity, immune characteristics, and the lack of ethical barriers to their procurement, make them ideal candidates for the basis of further research into their potential for disease treatment. Established methods for the isolation, cryopreservation, and culture of hAECs involve the use of animal products. Current regulations for the use of human cells as a cell therapy require the isolation process to be performed with animal product–free reagents, enzymes, and growth media (Therapeutic Goods Administration, 1989).

Critical Parameters and Troubleshooting Critical parameters we have encountered have been previously described by Miki et al. (UNIT 1E.3). This includes variability in cell number and cell viability between samples ob-

tained from different individuals, with significant decreases in these parameters following the storage of the placenta or amnion for >3 hr before isolation of hAECs. We found that the greatest factor influencing the yield of cells is contamination with blood cells. Thorough washing of the placenta and the amnion membrane is recommended as it appears that minor blood cell contamination is sufficient to inhibit the activity of the TrypZean solution. Contamination of the cell yield with mesenchymal cells can be avoided by decreasing the digest period. This will also decrease the total yield of cells. We found that EpiLife growth medium was a suitable animal product–free culture medium for the maintenance and expansion of hAECs. However, optimization of growth factor concentrations is required to achieve optimal growth rates for this cell type. In this study, we have addressed the issue of cell adherence during culture by using a human recombinant collagen coating matrix, which significantly increased the plating efficiency. However, following prolonged exposure to TrypZean the plating efficiency is still sub-optimal.

Anticipated Results The animal product–free method for the isolation of hAECs developed by our group results in a cell population that is comparable to the previously established animal product– containing method, producing an average yield of 120 ± 40 × 106 hAECs with an average viability of 83 ± 4% (Fig. 1E.6.1A). Isolates were shown to consist of 92% EpCAM positive cells with <1% mesenchymal cell contamination (Fig. 1E.6.1B,C). Analysis with a panel of monoclonal antibodies specific for cell surface antigens demonstrated consistency in the isolated cell populations from different individuals (data not shown). Interestingly, following the culture of hAECs for 5 passages, the cell surface marker profile of the population is significantly different. Table 1E.6.3 shows a selection of surface markers, which demonstrates this change. This should be taken into consideration if hAECs are to be maintained through serial passaging. There was no significant difference in post-thaw viability between serum-based and animal product–free cryopreservation media (Fig. 1E.6.3A). However, metabolic recovery, as determined by an MTS assay, was greatest when hAECs were cryopreserved in Cryostor cryopreservation medium (Fig. 1E.6.3B). We have previously administered hAECs into the testes of immune-compromised mice, with

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Table 1E.6.3 Human Amnion Epithelial Cells Interrogated by a Panel of Monoclonal Antibodies and Analyzed by Flow Cytometrya

Percent positive cells BD mAB Antibody

hAEC P0

hAEC P5

CD1b

5

88

CD10

80

0

CD26

72

10

CD29

94

98

CD34

6

97

CD45

1

9

CD46

55

97

CD55

100

98

CD58

5

99

CD59

100

69

CD63

98

0

CD66f

0

71

CD73

98

14

CD77

30

88

CD81

35

99

CD90

2

0

CD91

30

92

CD95

15

28

CD98

90

99

CD104

87

95

CD108

3

78

CD109

15

0

CD142

98

98

CD147

100

99

CD151

97

25

CD164

97

78

CD166

28

77

CD227

98

75

EGF-R

30

76

fII-R

99

86

HLA-ABC

93

41

HLA-A2

0

0

HLA-DQ

16

71

HLA-DR

0

0

a The numbers represent the percentage of the cells within each cell

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

type that are positive for the specific marker. Marker expression is shown for freshly isolated human amnion epithelial cells (P0) and after 5 passages (P5). The surface marker profile shown was selected due to either the differential expression by P0 and P5 hAECs, or the biological importance of the antigen.

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A

B

1

2

3

4

1,250

5

6

13

7

8

14

9

10

15

16

11

17

12

18

Count

1,000 750 500

S

250

G1

0 19

20

21

22

23

24

G2/M

50 100 150 200 250 Vybrant DyeCycle Violet fluorescence

C Telomere length (kbp)

20 15 10 5 0 BM MSC

ESC

hAEC PO hAEC PS

H control

L control

Figure 1E.6.4 Characterization of cultured hAECs. (A) Giemsa band karyogram showing chromosomes of passage 5 hAECs. (B) Flow cytometry of passage 5 hAECs showing DNA stained with DyeCycle Violet. G1 and G2/M indicate 2n and 4n cellular DNA content, respectively; S indicates cells undergoing DNA synthesis, intermediate in DNA content between 2n and 4n. (C) Telomere lengths of bone marrow–derived mesenchymal stromal cells (BM MSC), hESC, and hAECs when freshly isolated (P0) and at passage 5 (P5). High length control (H) and low length control (L) telomere standards are provided in the assay kit. Average telomere lengths are indicated (C).

no palpable tumors detected 10 weeks postinjection (Ilancheran et al., 2007). This lack of tumorigenicity is supported by the observation that hAECs display a normal karyotype and cell cycle distribution after 5 passages while maintaining long telomere lengths (Fig. 1E.6.4). These properties indicate that, unlike human embryonic stem cells (Trounson and Pera, 2001) and human induced pluripotent stem cells (Yu et al., 2007), hAECs are not likely to form tumors in clinical applications. Multipotent differentiation potential of hAECs, isolated using our animal product– free method, was demonstrated in vitro by the maintenance of hAECs in conditions known to induce the differentiation of stem cells into lung epithelial, osteogenic, adipogenic, and neural lineages. We demonstrated that culture in these conditions resulted in the induction of lung epithelial (SP-C, SP-D; Fig. 1E.6.5A), osteogenic (osteocalcin, osteonectin), adipogenic (PPARγ, LPL; Fig. 1E.6.5B), and neural (Nestin, β-3-Tubulin) specific gene

expression (Fig. 1E.6.5C). Culture of hAECs under these conditions also resulted in the induction of lung epithelial (Fig. 1E.6.5D), glial, and neural-specific protein expression (ProSP-C, GFAP and MAP2 for lung epithelium, glia, and neurons respectively; Fig. 1E.6.5F). Osteogenic and adipogenic differentiation was demonstrated by histological methods (Alzarin Red staining of mineralized calcium for osteogenic and Oil Red O staining of triglycerides for adipogenic differentiation; Fig. 1E.6.5E). We have now optimized an animal product–free method to allow efficient isolation and cryopreservation of hAECs for use in future clinical therapies.

Time Considerations

It takes ∼4 hr to isolate hAECs using Basic Protocol 1. Cryopreservation requires ∼20 min. Immunocharacterization of hAEC requires ∼3 hr. The cell proliferation assay requires 1 to 4 hr for completion of the Cell Titer 96 AQueous assay and ∼3 hr for the ELISA

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Differentiation of hAEC

A

B

Endoderm Day 0

Day 28 SAGM

Human lung

C

Mesoderm Day 0

NTC

Day 28

Day 28

Epilife

Diff

Ectoderm

NTC

SP-A

Osteocalcin

SP-B

Osteonectin

Nestin

SP-C

LPL

-3-Tubulin

SP-D

Day 0

PPAR

Day 28 Neural

Human brain

NTC

GAPDH

GAPDH

GAPDH

D

E Lung epithelium Pro-SP-C

F Osteocytes Alizarin Red

Adipocytes Oil-Red-O

Glial GFAP

Neural MAP2

Differentiation media

Negative control

Figure 1E.6.5 Multipotent differentiation potential of hAECs isolated with animal product–free method. Differentiation potential was demonstrated by the culture of hAECs in conditions developed to grow and maintain neurons, small airway epithelial cells, osteocytes, and adipocytes. PCR was used to detect expression of lung epithelial (A), osteogenic (B), adipogenic, glial, and neural (C) specific gene expression. Immunocytochemistry was performed to determine expression of Pro-SP-C (D) and GFAP and MAP2 (F). Alizarin Red staining was used to detect calcified deposition by cells, and Oil-Red-O staining was used to detect lipid in cells (E). Human brain and lung were used as negative and positive controls for PCR and immunocytochemistry.

assay. Assessing telomere length requires ∼18 hr.

Acknowledgments This study was funded by NH&MRC project grant no. 491145. The authors wish to thank Hayley Dickinson and Daniel Layton for their intellectual input, Natalie Seach for providing MSC genomic DNA, and Adam Goulburn for providing ESC genomic DNA. We also thank BD Biosciences for providing the monoclonal antibodies used in Support Protocol 1.

Literature Cited

Amnion Epithelial Cell Isolation and Characterization for Clinical Use

Bailo, M., Soncini, M., Vertua, E., Signoroni, P.B., Sanzone, S., Lombardi, G., Arienti, D., Calamani, F., Zatti, D., Paul, P., Albertini, A., Zorzi, F., Cavagnini, A., Candotti, F., Wengler, G.S., and Parolini, O. 2004. Engraftment potential of human amnion and chorion cells derived from term placenta. Transplantation 78:14391448.

Brown, T. 1999. Southern blotting. Curr. Protoc. Mol. Biol. 68:2.9.1-2.9.20. Fernandes, M., Sridhar, M.S., Sangwan, V.S., and Rao, G.N. 2005. Amniotic membrane transplantation for ocular surface reconstruction. Cornea 24:643-653. Hou, Y., Huang, Q., Liu, T., and Guo, L. 2008. Human amnion epithelial cells can be induced to differentiate into functional insulin-producing cells. Acta Biochim. Biophys. Sin. 40:830-839. Ilancheran, S., Michalska, A., Peh, G., Wallace, E.M., Pera, M., and Manuelpillai, U. 2007. Stem cells derived from human fetal membranes display multi-lineage differentiation potential. Biol. Reprod. 77:577-588. Kakishita, K., Elwan, M.A., Nakao, N., Itakura, T., and Sakuragawa, N. 2000. Human amniotic epithelial cells produce dopamine and survive after implantation into the striatum of a rat model of Parkinson’s disease: A potential source of donor for transplantation therapy. Exp. Neurol. 165:2734.

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Kakishita, K., Nakao, N., Sakuragawa, N., and Itakura, T. 2003. Implantation of human amniotic epithelial cells prevents the degeneration of nigral dopamine neurons in rats with 6-hydroxydopamine lesions. Brain Res. 980:4856. Laws, P.J. and Hilder, L. 2008. Australia’s mothers and babies 2006. Perinatal statistics series no. 22. Cat. no. PER 46. Sydney. AIHW National Perinatal Statistics Unit. Li, H., Niederkorn, J.Y., Neelam, S., Mayhew, E., Word, R.A., Mcculley, J.P., and Alizadeh, H. 2005. Immunosuppressive factors secreted by human amniotic epithelial cells. Invest. Ophthalmol. Vis. Sci. 46:900-907. Parolini, O., Alviano, F., Bagnara, G.P., Bilic, G., Buhring, H.J., Evangelista, M., Hennerbichler, S., Liu, B., Magatti, M., Mao, N., Miki, T., Marongiu, F., Nakajima, H., Nikaido, T., Portmann-Lanz, C.B., Sankar, V., Soncini, M., Stadler, G., Surbek, D., Takahashi, T.A., Redl, H., Sakuragawa, N., Wolbank, S., Zeisberger, S., Zisch, A., and Strom, S.C. 2008. Concise review: Isolation and characterization of cells from human term placenta: Outcome of the first international Workshop on Placenta Derived Stem Cells. Stem Cells 26:300-311. Reubinoff, B.E., Pera, M.F., Fong, C.-Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotech. 18:399-404.

Sankar, V. and Muthusamy, R. 2003. Role of human amniotic epithelial cell transplantation in spinal cord injury repair research. Neuroscience 118:11-17. Therapeutic Goods Act. 1989. C2009C00028. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Trelford, J.D. and Trelford-Sauder, M. 1979. The amnion in surgery, past and present. Am. J. Obstet. Gynecol. 134:833-845. Trounson, A. and Pera, M. 2001. Human embryonic stem cells. Fertility and Sterility 76:660-661. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Wei, J.P., Zhang, T.S., Kawa, S., Aizawa, T., Ota, M., Akaike, T., Kato, K., Konishi, I., and Nikaido, I. 2003. Human amnion-isolated cells normalize blood glucose in streptozotocininduced diabetic mice. Cell Transplant. 12:545552. Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S., Nie, J., Jonsdottir, G.A., Ruotti, V., Stewart, R., Slukvin, I.I., and Thomson, J.A. 2007. Induced pluripotent stem cell lines derived from human somatic cells. Science 318:19171920.

Embryonic and Extraembryonic Stem Cells

1E.6.25 Current Protocols in Stem Cell Biology

Supplement 13

Differentiation of Embryonic Stem Cells into Cartilage Cells

UNIT 1F.1

Jaspal Singh Khillan1 1

University Of Pittsburgh, Pittsburgh, Pennsylvania

ABSTRACT Embryonic stem (ES) cells have complete potential to form all types of cells. Although these cells have indefinite capacity for self-renewal, the mechanisms that control their lineage-restricted differentiation are not well understood. Due to their potential to form all types of cells, these cells are expected to have applications in regenerative medicine to cure human diseases. Osteoarthritis (OA) is a degenerative disease of articular cartilage of weight bearing joints. Approximately twenty million people suffer from this debilitating disease. Therefore, the induced differentiation of ES cells into cartilage-producing cells will have potential application to cure OA. This unit describes a system to induce differentiation of a high percentage of ES cells into mesenchymal cells that differentiate into chondrocytes, the cartilage-producing cells. A quantitative production of chondrocytes can be a powerful resource to alleviate the suffering of those patients with OA. Furthermore, this can be an excellent system to investigate the upstream events of cell-restricted differentiation during the inaccessible period of development. Curr. Protoc. Stem Cell C 2007 by John Wiley & Sons, Inc. Biol. 2:1F.1.1-1F.1.13.
Keywords: embryonic stem cells r limb bud progenitor cells r cartilage r chondrocytes r collagen type II r alternate splicing of collagen type II gene r Oct-4 transcription factor r proteoglycans

INTRODUCTION Embryonic stem (ES) cells derived from the inner cell mass (ICM) of the blastocyst have complete potential to form all the primary germ layers such as ectoderm, mesoderm, and endoderm. The cellular and molecular mechanisms that control their differentiation into specific lineages are not fully elucidated. After microinjection into blastocysts, these cells integrate and populate all the lineages including germ line cells; this has led to the assumption that ES cells undergo the same series of developmental changes during differentiation into various cell lineages as normal inner cell mass. Under the conditions that promote differentiation, the ES cells differentiate spontaneously into embryoid bodies (EBs), which contain many types of cells including neurons, insulin-producing cells, bone-forming cells, and hematopoietic cells. Because EBs contain cells that are committed to different lineages, they are not suitable for cell fate determination. Recent developments in human ES cells have offered a challenge to develop strategies for understanding the basic mechanisms that play key roles in lineage-restricted differentiation for their applications in regenerative medicine and cell-based therapies. This protocol describes a micromass culture system for induced differentiation of ES cells into mesenchymal cells for quantitative production of cartilage-producing cells such as chondrocytes. Further this system may also be a useful model to investigate the upstream events of lineage-restricted differentiation of stem cells. The unit describes a co-culture system for ES cells with the limb bud progenitor cells (Basic Protocol), the preparation of feeder cells by mitomycin treatment and γ irradiation (Support Protocol 1), and preparation of limb bud progenitor cells (Support Protocol 2). In addition protocols are described for Alcian blue staining of differentiated cells Current Protocols in Stem Cell Biology 1F.1.1-1F.1.13 Published online July 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f01s2 C 2007 John Wiley & Sons, Inc. Copyright


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Figure 1F.1.1 ES cells after 48 hr of culture. Individual colonies grow over a monolayer of feeder cells. Each colony contains about 200 to 300 cells and is recognizable by distinct shiny edges. (100× magnification)

(Support Protocol 3), analysis of differentiated cells by RT-PCR (Support Protocol 4), and characterization of gene expression (Support Protocol 5). NOTE: The experiments described here require tissue culture facilities. Tissue culture and media preparation should be performed under sterile conditions using laminar flow hood. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals. BASIC PROTOCOL

PREPARATION, CO-CULTURE, AND ANALYSIS OF DIFFERENTIATED EMBRYONIC STEM CELLS ES cells are cultured in ES medium over mitotically inactive fibroblasts feeders in a humidified incubator with 5% CO2 at 37◦ C, following methods previously described (Robertson, 1997). Figure 1F.1.1 shows typical colonies of ES cells after 48 hr. The distinct individual phase-bright colonies of ES cells can be seen scattered throughout the plate. The ES medium is supplemented with 1000 IU/ml of leukemia inhibitory factor (LIF) to prevent the differentiation of ES cells. Pluripotent mouse ES cells require a lymphokine, LIF, to maintain their self-renewal. The feeder cells support undifferentiated ES cells by providing LIF; however, the culture medium should also be supplemented with recombinant LIF to prevent differentiation. To induce differentiation, ES cells are co-cultured with 25% limb bud progenitor cells (LBPC) in high-density (10 to 20 × 106 cells/ml) micromass cultures (Ahrens et al., 1977). NOTE: All the cell cultures are maintained in humidified incubators with 5% CO2 at 37◦ C temperature. The medium should be warmed to room temperature before adding to the cells.

Materials

Differentiation of Embryonic Stem Cells into Cartilage Cells

Gelatin solution (see recipe) 1.0 × 106 mitomycin C–treated or irradiated primary mouse fibroblast cells (Support Protocol 1) ES medium (see recipe) Frozen ES cells 0.25% (w/v) trypsin/EDTA (Invitrogen)

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PF medium (see recipe) Calcium- and magnesium-free phosphate-buffered saline (CMF-PBS: Invitrogen) Plasmid DNA for neo and/or enhanced green florescence protein (EGFP; e.g., pEGFPN1; Clontech) 100-mm plates with feeder cells G418 (Gibco/BRL) 10% (v/v) DMSO in ES medium LBPC cells (Support Protocol 2) Trypsin 2 mg/ml collagenase 60-mm tissue culture dishes Electroporation vials Bio-Rad Pulsar II electroporator 96-well plate 24-well feeder coated plate Stereomicroscope with UV attachment Rotator Four-well Nunc Petri dishes (Fisher Scientific) Microcentrifuge tubes Additional reagents and equipment for preparation of mitomycin C–treated fibroblast cells (Support Protocol 1), cell counting (Phelan, 2006), PCR (Kramer and Coen, 2001), and preparation of limb bud progenitor cells (LBPC; Support Protocol 2) Prepare feeder cells 1. Coat a 60-mm plate by incubating with 2 to 3 ml of gelatin solution for at least 2 hr. 2. Transfer 1.0 × 106 mitomycin C–treated or γ-irradiated primary mouse fibroblast cells to the gelatin-coated dish. Feeder cells are fibroblast cells that are mitotically inactivated before preparing plates. The cells are treated with 10 µg/ml mitomycin C for 2 hr. The cells are washed extensively in primary fibroblast (PF) culture medium and are used directly to prepare feeder plates. Alternatively the cells are irradiated at 4000 to 6000 rads, and divided into 1.0 × 106 cells/aliquot and stored in liquid nitrogen. (See Support Protocol 1)

3. Allow cells to settle overnight and change to ES medium the next day.

Culture ES cells 4. Thaw 1 vial of frozen ES cells in 37◦ C water bath for 90 sec. Transfer cells to 5 ml ES medium in a 15-ml tube to wash the cells. Centrifuge for 5 min at 200 × g, 20◦ C. Resuspend the pellet in 6 ml of ES medium and transfer all of the contents to a gelatinized plate. Several different ES cell lines are available from different laboratories including commercial sources. R1 cells were obtained from Dr. A. Nagy. ES cells can be maintained indefinitely under specialized culture conditions such as growth over feeder cells (UNIT 1C.1) and supplementing medium with LIF. In general ES cells require about 2000 IU of LIF/ml. The feeder cells provide about 1000 IU, the rest is provided by supplementation.

5. Incubate, changing the medium every 24 hr until the cells are confluent (∼48 to 72 hr). For ES cell culture, the medium must be changed every day and the cells must be passaged every 48 to 72 hr, otherwise the cells will start to differentiate. The cells must be trypsinized before the colonies become too large. After trypsinization the feeder cells can be separated from ES cells by letting the suspension stand. Feeder cells settle down faster because they are heavier than the ES cells leaving pure ES cells on the top. The cells at the bottom also contain many ES cells which can be saved for future use. Current Protocols in Stem Cell Biology

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Passage ES cells 6. Trypsinize cells with 1 ml 0.25% trypsin/EDTA for 5 min and neutralize trypsin by adding 5 ml of ES medium. 7. To disperse cells to a single-cell suspension pipet 10 to 15 times and transfer cells into a 15-ml tube. Centrifuge cells 5 min at ∼200 × g, 20◦ C. 8. Resuspend the cells in 10 ml PF medium. 9. Place cells on ice for 10 min to allow feeder cells to settle. 10. Transfer top 8 ml of the cell suspension to a new 15-ml tube. Perform another round of 10 min sedimentation (step 9) and collect top 6 ml of feeder-free cell suspension. 11. Centrifuge cells for 5 min at 200 × g, 20◦ C and wash cells at least two times in 5 ml PF medium followed by resuspension in 5 ml of fresh PF medium. Check cell concentration with a hemacytometer (Phelan, 2006).

Electroporate DNA 12. Transfer 1.0 to 2.0 × 106 ES cells into a 15-ml tube and centrifuge 5 min at 200 × g, 20◦ C . 13. Wash pellet in 5 ml CMF-PBS once and resuspend cells in 700 µl of CMF-PBS. 14. Dissolve 25 to 30 µg plasmid DNA for neo and/or EGFP in 100 µl of CMF-PBS. ES cells are prepared to contain selectable gene markers such as neomycin phospho transferase (neo) or EGFP. The cells are electroporated with pEGFPN1 plasmid (Clonetech), which contains neo and enhanced green florescence protein (EGFP) gene. The electroporated cells are selected with 150 to 250 µg/ml G418 and individual clones are collected. The EGFP can be used as an additional marker to select colonies under UV microscope.

15. Mix DNA with ES cells and transfer all of the contents to an electroporation vial. 16. Keep cells on ice for 10 min. 17. Electroporate DNA using Bio-Rad Pulsar II at 250 V and 500 µF.

Plate and select cells 18. Allow cells to stand at room temperature for 5 min and transfer equal amounts onto four 100-mm plates with feeder cells. 19. Add 12 ml ES medium and allow cells to settle overnight. 20. After 24 hr add 12 ml fresh ES medium with 150 to 250 µg/ml G418. Return cells to incubator. Neo-resistant individual colonies of ES cells are formed after 8 to 10 days

Expand colonies 21. Scrape colonies individually and transfer to 96-well plate with 50 µl 0.25% trypsin/EDTA (Invitrogen). 22. After 4 min add 100 µl ES medium and transfer contents into a 24-well feeder-coated plate 23. Collect ∼40 to 50 colonies in each individual well of a 24-well plate and culture for about 3 to 4 days until the cells are confluent. Differentiation of Embryonic Stem Cells into Cartilage Cells

24. Trypsinize cells (step 6) and freeze half the cells in 10% DMSO in ES medium and use other half for DNA analysis.

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25. Check the presence of neo gene by PCR (Kramer and Coen, 2001) or by EGFP florescence using a UV microscope. Use Southern blot (Brown, 1999) analysis to confirm the integrity of the DNA.

Co-culture ES cells and progenitor cells 26. Transfer 1.5 × 105 ES cells and 50,000 LBPC cells in 500 µl of PF medium to a microcentrifuge tube. 27. Mix cells on a rotator for 15 min (20 to 30 revolutions per min) and follow with a 15 min incubation at 37◦ C. 28. Repeat step 27 two times, and then pellet cells in a microcentrifuge 5 min at 200 × g, 20◦ C. 29. Resuspend cells in 20 to 25 µl of PF medium and transfer total contents to a 4-well Nunc plate, using the same number of ES cells and LBPC in separate wells as controls. 30. Establish parallel cultures of pure ES cells and LBPC separately as negative and positive controls respectively. 31. Add ∼800 µl of PF medium after 2 hr to cover the cells. 32. After overnight culture, examine the cultures. ES cells form typical multicellular colonies (Fig. 1F.1.2B), whereas co-cultured cells show appearance of a sheet of flat cells (Fig. 1F.1.2A).

33. Return the cells to the incubator. Change ∼1 ml PF medium every other day. 34. After 4 days, treat cells with 100 µg/ml G418 to select against LBPC. 35. Monitor differentiation of cells by the appearance of aggregates of cells (Figure 1F.1.3A, LBPC and Figure 1F.1.3B, ES cells). The pure mesenchymal cells from limb buds form aggregates or nodules of differentiating cells after 3 to 4 days of culture. On the other hand, condensations in induced ES cells appear after about 7 days, which suggests that first 3 to 4 days are required for induction of ES cells into chondrogenic lineage.

36. Harvest cells at different time intervals (such as 3 days and 7 days) for protein and RNA analyses. Scrape the cell aggregates and surrounding cells and dissociate by treating with trypsin and 2 mg/ml collagenase. Count cells with a hemacytometer (Phelan, 2006).

Figure 1F.1.2 Co-culture of ES cells with LBPC. (A) ES cells were co-cultured with 25% LBPC in high-density micromass culture. The cells form a uniform flat layer of cells after 24 to 48 hr as compared to (B) pure ES cells that form only multicellular colonies typical of undifferentiated embryonic stem cells. (Magnification, 100×).

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Figure 1F.1.3 Differentiation of ES cells into chondrocytes. Micromass cultures were established separately for (A) pure limb bud mesenchymal cells and (B) LBPC and ES cell co-cultures. Pure LBPC form aggregates of condensed cells within 3 to 4 days. These aggregates develop into nodules (marked by arrows) of differentiated cells after 10 to 12 days of culture (A). Each nodule is usually surrounded by a unicellular layer of perichondrial cells. After 4 days of co-culture, the cells were treated with G418 to remove the LBPCs. The remaining cells formed aggregates after 7 to 8 days (∼3 to 4 days after that seen in pure LBPC cultures) that develop into nodules of chondrocytes at 14 to 15 day (B). Unlike pure LBPC, the nodules in mixed cultures are surrounded by a small number of indistinct cells (dark looking cells). These cells represent ES cells that failed to differentiate into chondrocytes. Cultures of pure ES cells did not form nodules (not shown; magnification, 100×).

SUPPORT PROTOCOL 1

PREPARATION OF MITOTICALLY INACTIVE FEEDER CELLS Confluent cultures of fibroblasts are established on 100-mm plates. The cells are treated with 10 µg/ml mitomycin C solution (see recipe) for 2 hr. The cells are then washed extensively in PF culture medium to remove mitomycin C. These cells can be used directly to prepare feeder plates. Alternatively, the trypsinized cells can be irradiated at 4000 to 10000 rads. The cells are then washed and stored in liquid nitrogen as small aliquots of 1.0 × 106 cells per vial. Under these conditions the cells can be stored indefinitely.

SUPPORT PROTOCOL 2

PREPARATION OF LIMB BUD PROGENITOR CELLS (LBPC) The progenitor cells are obtained from limb buds of E11.5 embryos. Adult 5- to 6-weekold FVB/N females are mated with normal 10- to 12-week-old stud males. To obtain E11.5 embryos, the females are mated with the stud males. Usually 2 to 3 females are mated with each male late in the evening. The pregnant females are checked the next morning for the presence of cream-colored vaginal plugs formed due to the hardening of secretions in the semen. The day of mating is considered E0.5.

Materials Breeding pairs of FVB/N (Taconic Farm) mice consisting of: Females: 5- to 6-week old Males: 10- to 12-week old stud males CO2 CMF-PBS 0.25% (w/v) trypsin/EDTA (Invitrogen) PF medium (see recipe)

Differentiation of Embryonic Stem Cells into Cartilage Cells

100-mm petri dishes Surgical instruments, e.g., two pairs of forceps and a pair of scissors for collection of embryos Additional reagents and equipment for euthanizing mice by CO2 asphyxiation (Donovan and Brown, 2006)

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Figure 1F.1.4 Limb buds isolated from E11.5 embryos. To prepare limb bud progenitor cells (LBPC), limb buds from several embryos are pooled.

Set up breeding pairs 1. Set up mating of 2 to 3 breeding pairs of mice. 2. Verify mating by the presence of vaginal plugs the following morning. The day of plug is E0.5.

3. Sacrifice pregnant females on day 11 by CO2 asphyxiation (Donovan and Brown, 2006). The embryos will be approximately at E11.0 to E11.5.

Collect limb bud progenitor cells 4. Collect uterine horns from the mothers and place them in 100-mm petri dish with CMF-PBS. 5. Using scissors, cut uterine wall and dispense embryos in a fresh petri dish with 10 ml CMF-PBS. 6. Wash embryos extensively with CMF-PBS to remove blood using a fresh petri dish each time. 7. Excise limb buds from each embryo (Fig. 1F.1.4). 8. Trypsinize limb buds from 8 to 10 embryos in 0.25% trypsin/EDTA for 5 to 6 min. 9. Triturate the tissue in 10 ml of PF medium to a single-cell suspension. 10. Wash cells by centrifuging 5 min at 200 × g, 20◦ C, two times in 10 ml PF medium and resuspend the final pellet in 10 ml of the same medium in a 15-ml tube. 11. Allow cells to stand for 3 to 5 min for tissue pieces to settle down. 12. Transfer top 8 ml into a fresh 15-ml tube and pellet cells by centrifuging 5 min at 200 × g, 20◦ C. 13. Resuspend pellet in 5 ml of PF medium and check cell concentration with a hemacytometer (Phelan, 2006) and store cells in a 37◦ C incubator until further use.

ALCIAN BLUE STAINING OF DIFFERENTIATED CELLS Terminally differentiated chondrocytes express cartilage-specific sulfated proteoglycans that stain positive with the Alcian blue. The differentiated chondrocytes, therefore, can be stained with Alcian blue after 14 days.

SUPPORT PROTOCOL 3

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Supplement 2

Figure 1F.1.5 Alcian blue staining of differentiated cells. ES cells were co-cultured with LBPC followed by G418 treatment after 4 days. At day 19, the cells were stained with Alcian blue. In general the nodules formed by differentiated ES cells exhibit diffused boundaries as compared to normal LBPC. (B) Densely packed and overlapping nodules formed by the differentiated cells that stained positive with the dye. Staining of pure LBPC cultures show individual nodules stained with the dye (A). No Alcian blue staining was observed in parallel culture of pure ES cells (not shown; magnification, 100×).

Materials Micromass cultures containing putative chondrocytes (Basic Protocol) CMF-PBS 100% ethanol Alcian blue dye (see recipe) 80% (v/v) glycerol with distilled H2 O 100% glycerol Additional reagents and equipment for micromass cultures containing putative chondrocytes (Basic Protocol) 1. Wash micromass cultures (Basic Protocol) containing putative chondrocytes two times with 1 ml PBS. 2. Fix cells in 1 ml 100% ethanol for 1 hr. 3. Stain cells for 2 to 4 hr with 1 ml Alcian blue dye. 4. Wash cells several times with 1 ml 100% ethanol to remove excess dye. 5. Clarify cells with 80% glycerol solution (Fig. 1F.1.5). 6. Store samples indefinitely under 100% glycerol. 7. Score cultures for blue-stained nodules (Fig. 1F.1.5) SUPPORT PROTOCOL 4

Differentiation of Embryonic Stem Cells into Cartilage Cells

ANALYSIS OF DIFFERENTIATED CELLS BY REVERSE TRANSCRIPTASE–POLYMERASE CHAIN REACTION (RT-PCR) Differentiated cells are analyzed for expression of cartilage cell–specific genes. G418treated micromass cultures are harvested at different intervals by trypsinization and total RNA is isolated. RT-PCR analysis is carried out using primers specific for marker genes such as POU domain containing Oct-4 for undifferentiated cells (Fig. 1F.1.6) and collagen type II for differentiated cells. Collagen type II is the most abundant protein in the extracellular matrix of cartilage; it is expressed in two forms due to alternate splicing of the exon 2 of the message. The prechondrogenic mesenchymal cells express exclusively the unspliced form type IIA, whereas mature chondrocytes contain shorter transcript, type IIB, in which exon 2 is spliced out. The alternate spliced forms are identifiable by RT-PCR analysis using specific primers that amplify 489-bp and 285-bp fragments respectively (Fig. 1F.1.7).

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Current Protocols in Stem Cell Biology

Figure 1F.1.6 Expression of undifferentiated cell–specific transcription factor Oct-4. Undifferentiated ES cells express transcription factor Oct-4 that disappears as the cells differentiate. RT-PCR analysis shows that undifferentiated ES cells express transcription factor Oct-4 (lane 1). After 7 days, the expression of Oct-4 decreased dramatically (lane 2), which disappeared completely in differentiated cells at day 19 (lane 3). LBPC at day 1 do not express Oct-4 (lane 4).

Figure 1F.1.7 Expression of chondrocytespecific genes. ES cells co-cultured with 25% LBPC were treated with G418 after 4 days. The cells were washed to remove dead cells and harvested at day 7 followed by isolation of total RNA. Lanes 1 to 3 show RT-PCR analysis for collagen type II. Lane 1, RNA from pure LBPC at day 1; lane 2, RNA from LBPC at day 7; lane 3, RNA from co-cultured and G418-treated cells at day 7. As expected, pure LBPC on day of isolation amplify only type IIA-specific 489-bp fragment (Lane 1) whereas, the same cells at day 7 amplified DNA fragments for type IIA (489-bp) and type IIB (285-bp) transcripts representing cells differentiated into chondrocytes (lane 2). On the other hand, ES cells co-cultured with LBPCs amplified only 489bp fragment specific for type IIA transcript (lane 3) similar to that seen in day 1 prechondrogenic cells. No such amplification was seen in pure ES cell cultures (not shown).

Materials G418-treated micromass cultures (Basic Protocol) 0.25% (w/v) trypsin/EDTA RNA extraction kit (STAF-60; Tel-Test) RT-PCR kit (Invitrogen) Tris KCl MgCl2 Taq DNA polymerase PCR primers: Oct-4:(forward) 5 -GGCGTTCTCTTTGGAAAGGTGTTC-3 (reverse) 5 -CTCGAACCACATCCTTCTCT-3 Collagen II:(forward) 5 -GTGAGCCATGATCCGC-3 (reverse) 5 -GACCAGGATTTCCAGG-3 (Carlberg et al., 2001) Neomycin:(forward) 5 -AGGATCTCCTGTCATCTCACCTTGCTCCTG-3 (reverse) 5 - AAGAACTCGTCAAGAAGGCGATAGAAGGCG-3 HPRT:(forward) 5 -GTAATGATCAGTCAACGGGGGAC-3 (reverse) 5 -CCAGCAAGCTTGCAACCTTAACCA-3 2% (w/v) agarose gel Thermal cycler Additional reagents and equipment for RNA extraction (Kingston et al., 1996), RT-PCR (Beverly, 2001), and gel electrophoresis (Voytas, 2000) Current Protocols in Stem Cell Biology

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1. Harvest G418-treated micromass cultures at 3 days and 7 days using 0.25% trypsin/EDTA. 2. Extract the RNA using STAT-60. 3. Reverse transcribe the RNA using kit from Invitrogen. 4. Prepare polymerase chain reaction mix (50 µl per reaction).

100 mM Tris·Cl, pH 8.3 500 mM KCl 15 mM MgCl2 0.25 U Taq DNA polymerase 200 mM of each primer. 5. Using the appropriate primers, amplify Oct-4, collage II, neomycin, and HPRT using the following program: 35 to 40 cycles:

30 sec 30 sec 90 sec

94◦ C 60◦ C 72◦ C

(denaturation) (annealing) (extension).

Primers for hypoxanthine phosphoribosyl transferase (HPRT) are used as control.

6. Separate the amplified fragments on 2% agarose gels (Voytas, 2000). The expression of 489-bp fragment in micromass cultures is seen at ∼7 days suggesting that first 3 to 4 days are sufficient for the induction of ES cells into a stage equivalent to prechondrogenic cells.

SUPPORT PROTOCOL 5

DIFFERENTIAL GENE EXPRESSION ANALYSIS For analyses of differentially expressed genes during cell fate determination, GFPpositive ES cells are mixed with normal LBPC. After thorough mixing, the cells are plated as micromass cultures. After different intervals of time, the cells are trypsinized and separated by fluorescent activated cell sorter for GFP-positive cells. The induced ES cells can be used directly for RNA isolation for further stages of chondrocyte differentiation. The short form of collagen mRNA appears as soon as the cells form nodules, which is usually in 8 to 10 days.

Materials 1.5 × 106 GFP-positive ES cells (from frozen stocks) LPBC from normal embryos (Support Protocol 2) PF medium (see recipe) 4-well plate FACS (fluorescence-activated cell sorter) for GFP Additional reagents and equipment for preparation of LPBC (Support Protocol 2) 1. Mix 1.5 × 106 GFP-positive ES cells with 500,000 LBPC (25%) from normal embryos. 2. After thorough mixing, centrifuge cells for 5 min at 200 × g, 20◦ C and resuspend cells in 100 µl of PF medium. Differentiation of Embryonic Stem Cells into Cartilage Cells

3. Transfer 25 µl of cell suspension to each well in 4-well plate. Incubate cells at 37◦ C for 2 hr and following incubation add 800 µl of the medium.

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4. Change medium every other day using 1 ml PF medium. 5. Harvest cells at specific intervals such as 1, 2, 3 and 4 day by trypsinization using 0.25% trypsin/EDTA. 6. Separate induced ES cells in a fluorescence-activated cell sorter. The pure mesenchymal cells from limb buds form aggregates or nodules of differentiating cells after 3 to 4 days of culture. On the other hand, condensations in induced ES cells appear after about 7 days, which suggests that the first 3 to 4 days are required for induction of ES cells into chondrogenic lineage

7. Use cells directly for RNA isolation and differential gene expression by microarray analysis.

REAGENTS AND SOLUTIONS For culture recipes and steps, use Milli-Q purified water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Alcian blue dye solution 20 mg Alcian blue powder (Sigma; no. A3157) Dissolve in 80 ml ethanol and 20 ml glacial acetic acid (Kimmel and Trammell, 1981) Filter using Whatman-1 filter paper Make solution fresh prior to use The solution is filtered to remove dye particles.

Embryonic stem cell (ES) medium 90 ml of FBS 6 ml 10 mM non-essential amino acids (Invitrogen; no. 11140-050) 6 ml 100× penicillin/streptomycin 1 ml 2-mercaptoethanol (Invitrogen; no. 21985-023) 60 µl of 1000 U/ml leukemia inhibitory factor (LIF; Chemicon International) Store up to 4 weeks at 4◦ C (protected from light) Gelatin solution Dissolve 1.0 g gelatin (Sigma; no. 6650) in 1 liter deionized water. Store indefinitely in 50 to 100 ml aliquots at room temperature.

Mitomycin C solution Prepare a stock solution of mitomycin C 0.5 to 1.0 mg/ml (Sigma; no. M5030) in CMF-PBS or sterilized water. Store up to 1 week at 4◦ C protected from light.

Primary fibroblast (PF) culture medium Combine 10% (v/v) fetal bovine serum (FBS; Gemini-Bioproducts) and 1% penicillin/streptomycin (Invitrogen; no.15140-163) in Dulbecco’s Modified Eagle’s medium (DMEM; Invitrogen, no. 12430-054). Store up to 4 weeks at 4◦ C.

COMMENTARY Background Information Embryonic stem (ES) cells derived from the inner cell mass (ICM) of the blastocyst have complete potential to form all the primary germ layers such as ectoderm, mesoderm, and endoderm (Evans and Kaufman,

1981; Martin, 1981); however, the cellular and molecular mechanisms that control differentiation into specific lineages are not fully elucidated. Mouse ES cells can be maintained indefinitely without differentiation in the presence of a cytokine leukemia inhibitory factor

Embryonic and Extraembryonic Stem Cells

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Differentiation of Embryonic Stem Cells into Cartilage Cells

(LIF; Robertson, 1997). After microinjection into blastocyst, these cells integrate and populate all the lineages including germ line cells, e.g., ovary and testis (Bradley et al., 1984). These observations have led to the assumption that ES cells undergo the same series of developmental changes such as differentiation into various cell lineages and tissue development as normal ICM. Despite this potential, only limited information is available about the mechanisms responsible for the commitment of ES cells to the specific cell types. In the absence of LIF, ES cells differentiate spontaneously into embryoid bodies (EB; Doetschman et al., 1985) that contain many types of cells including neurons, chondrocytes, insulin-producing cells, bone-forming cells, and hematopoietic cells (O’Shea, 2001). Because EBs contain cells committed to many different lineages, they are not suitable for studies on cell fate determination. Recent developments with human ES cells have offered a challenge to develop strategies for understanding the basic mechanisms that play key roles in lineage-restricted differentiation of ES cells for their applications in regenerative medicine and cell-based therapies. This unit describes a micromass culture system for induced differentiation of ES cells into mesenchymal cells, which may be a useful model system to investigate the upstream events of stem cell differentiation into boneand cartilage-forming cells, i.e., osteoblasts and chondrocytes respectively. Injury or degeneration of cartilage in the joints is associated with osteoarthritis (OA) that affects the quality of life of over twenty million people in the United States of America. The quantitative production of chondrocytes therefore, can be a powerful resource to alleviate the suffering of those patients. This unit describes how ES cells can be induced to differentiate into chondrocytes by coculture with the mesenchymal progenitor cells isolated from limb buds of developing embryos. The ES cells and the progenitor cells are cultured in high-density micromass cultures to establish maximum contact between two types of cells which appear to transduce signals for the differentiation. The induction of differentiation is achieved within 3 to 4 days of contact after which the progenitor cells are removed either by their sensitivity to a drug such as the neomycin analog G418 or by fluorescenceactivated cell sorting (FACS) using a florescent marker gene such as GFP. The differentiated cells exhibit typical morphological characteristics of chondrocytes and express cartilage

matrix genes such as collagen type II and proteoglycans. No additional growth factors are required suggesting that signals from the progenitor cells are sufficient to induce ES cells into the chondrogenic lineage. Collagen type II is the most abundant protein in the extracellular matrix of cartilage. It is expressed in two forms due to alternate splicing of exon 2 of the mRNA (Sandell et al., 1994). The prechondrogenic mesenchymal cells express exclusively the unspliced form i.e., type IIA transcript, whereas mature chondrocytes contain type IIB transcript in which exon 2 is spliced out. Co-cultured ES cells and LBPC are treated with 100 µg/ml G418 at day 4 to eliminate limb bud–derived cells. After 3 days of treatment, i.e., after 7 days of culture, total RNA is tested by RT-PCR analysis using primers specific for type IIA (489 bp) and type IIB (285 bp) transcripts (Fig. 1F.1.7). Almost 60% of the ES cells differentiate into mature chondrocytes. The micromass culture therefore, can be an excellent model to study the upstream mechanisms of linage-restricted differentiation of stem cells and for quantitative production of chondrocytes.

Critical Parameters All tissue culture techniques should be performed under aseptic conditions in a Class II biohazard hood. The media must be sterilized by filtering through 0.22-µm filter. The reagents can be sterilized by autoclaving. Isolation of limb buds should be carried out under sterile conditions. The mice should be sacrificed by IACUC-approved protocols. Before collecting embryos, the animals should be sprayed with 70% ethanol. The PBS for washing the embryos and tissues should contain penicillin and streptomycin. ES cells and LBPC must be mixed thoroughly before co-culture. For mixing, the cells should be pipetted gently to prevent damage to the cells. After long cultures the differentiated cells tend to detach from the petri dish and should be handled carefully; therefore, add solutions very gently. The Alcian blue solution should be prepared fresh and should be filtered to remove dye particles.

Troubleshooting ES cells are sensitive to medium conditions. Make sure that the plates contain sufficient feeder cells. Add extra cells when required. If the medium appears too yellow, change to fresh medium quickly.

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Anticipated Results Normally, each limb bud can provide 25,000 to 50,000 cells. Usually, fifteen to twenty limb buds are sufficient for the experiment. After 7 days of co-culture, the ES cells begin to form nodules which become prominent after 10 days. Approximately 50% to 60% of cells differentiate into chondrocytes, as observed by Alcian blue staining. Compared to normal LBPC cultures, the nodules of differentiated ES cells are irregularly shaped and overlapping. Since the GFP is driven by CMV promoter, the cells exhibit GFP fluorescence throughout the culture, although it is slightly diminished after prolonged cultures.

Time Considerations Feeder cells Mitomycin treatment of mouse embryo fibroblasts takes ∼3 to 4 hr. Bulk γ-irradiated feeder cells can be prepared and frozen as aliquots of 1.0 × 106 cells. Each time a vial can be thawed for ES culture. Thawing of cells requires about 20 to 30 min. Embryonic stem cells Preparation of feeder-coated plates takes ∼2.5 to 3.0 hr including gelatinization of the plates. Thawing of ES cells takes ∼30 min. Confluent ES cells can be passaged in ∼30 min. Limb bud progenitor cells Isolation of embryos and collection of limb buds takes ∼1 hr. Mincing and trypsinization takes ∼30 min. Sedimentation and collection of single-cell preparation of LBPC takes ∼15 min. Co-culture of cells Mixing of cells requires ∼2 hr and setting up the micromass cultures take ∼30 min. Micromass cultures should be characterized at ∼7 days. Staining of cells Total time required for fixing the cells, staining, and clarification takes ∼7 to 8 hr.

Literature Cited Ahrens, P.B., Solursh, M., and Meier, S. 1977. The synthesis and localization of glycosaminoglycans in striated muscle differentiating in cell culture. J. Exp. Zool. 202:375-388.

Beverly, S.M. 2001. Enzymatic amplification of RNA by PCR (RT-PCR). Curr. Protoc. Mol. Biol. 56:15.5.1-15.5.6. Bradley, A., Evans, M., Kaufman, M.H., and Robertson, E. 1984. Formation of germ-line chimaeras from embryo-derived teratocarcinoma cell lines. Nature 309:255-256. Brown, T. 1999. Southern blotting. Curr. Protoc. Mol. Biol. 68:2.9.1-2.9.20. Carlberg, A.L., Pucci, B., Rallapalli, R., Tuan, R.S., and Hall, D.J. 2001. Efficient chondrogenic differentiation of mesenchymal cells in micromass culture by retroviral gene transfer of BMP-2. Differentiation 67:128-138. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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. Morph. 87:27-45. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156. Kimmel, C.A. and Trammell, C.A. 1981. A rapid procedure for routine double staining of cartilage and bone in fetal and adult animals. Stain Technol. 6:271-273. Kingston, R.E., Chomczynski, P., and Sacchi, N. 1996. Guanidine methods for total RNA preparation. Curr. Protoc. Mol. Biol. 36:4.2.1-4.2.9. Kramer, M.F. and Coen, D.M. 2001. Enzymatic amplification of DNA by PCR: Standard procedures and optimization. Curr. Protoc. Mol. Biol. 56:15.1.1-15.1.14. Martin, G.R. 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. 78:76347638. O’Shea, K.S. 2001. Directed differentiation of embryonic stem cells: Genetic and epigenetic methods. Wound Repair Regen. 9:443-459. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A3F.1-A.3F.18. Robertson, E.J. 1997. Derivation and maintenance of embryonic stem cell cultures. Methods Mol. Biol. 75:173-184. Sandell, L.J., Nalin, A.M., and Reife, R.A. 1994. Alternative splice form of type II procollagen mRNA (IIA) is predominant in skeletal precursors and non-cartilaginous tissues during early mouse development. Dev. Dyn. 199:129-140. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9.

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Differentiation of Human Embryonic Stem Cells to Cardiomyocytes by Coculture with Endoderm in Serum-Free Medium

UNIT 1F.2

Christine L. Mummery,1 Dorien Ward,1 and Robert Passier1 1

University of Utrecht Medical Centre, Utrecht, The Netherlands

ABSTRACT Many of the applications envisaged for human embryonic stem cells (hESC) undergoing cardiomyogenesis require that the differentiation procedure is robust and high yield. For many hESC lines currently available this is a challenge; beating areas are often obtained but subsequent analysis shows only few (<1%) cardiomyocytes actually present. Here the authors provide a protocol based on serum-free coculture with a mouse endoderm-like cell line (END2), which yields cultures containing on average 25% cardiomyocytes for two widely available hESC lines, hES2 and hES3. The authors also provide a variant on the protocol based on growth of hESC aggregates/embryoid bodies in END2-conditioned medium and a method for dissociating beating aggregates without compromising cardiomyocyte viability so that they can be used for transplantation into animals or further C 2007 (electrophysiological) analysis. Curr. Protoc. Stem Cell Biol. 2:1F.2.1-1F.2.14.
by John Wiley & Sons, Inc. Keywords: cardiomyocytes r human embryonic stem cells r coculture r hES2 r hES3 r visceral endoderm

INTRODUCTION Human embryonic stem cells (hESC) often differentiate spontaneously into multiple cell types when cultured as aggregates in suspension. These may then resemble embryoid bodies (EBs) that form when mouse embryonic stem cells are grown in suspension. These structures contain an inner core of epiblast-like cells and an outer endoderm “rind” and usually start to beat between 3 and 5 days after the aggregate has formed. Some hESC lines behave in a similar way and begin to beat in culture although this generally happens much later, at least 10 to 12 days after the initial aggregation step. The number of cardiomyocytes in these beating structures are generally low; in reports where numbers have been determined, not usually more than 1% to 2% of the cells within the beating aggregates are cardiomyocytes. Yet other lines do not differentiate spontaneously to cardiomyocytes at all. In this unit, a protocol is described which is effective in multiple cell lines passaged mechanically using the cut-and-paste method described in UNIT 1C.1 It is based on coculture with an endoderm-like cell line (END2) originally derived clonally from a differentiated population of P19 embryonal carcinoma stem cells (see Basic Protocol 1). It has also been described as effective in some primate ES cell lines, and END2-conditioned medium enhances cardiomyogenesis in hESC lines passaged enzymatically (UNIT 1C.1) if they are able to form EB-like structures in suspension (see Alternate Protocol). The protocols are designed on the assumption that the operator is familiar with mechanical and enzymatic passage of hESC and that the cells are maintained routinely on mouse or human feeder cells (UNIT 1C.1). The protocols describe the routine passage of END2 Mesodermal Lineages Current Protocols in Stem Cell Biology 1F.2.1-1F.2.14 Published online July 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f02s2 C 2007 John Wiley & Sons, Inc. Copyright


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cells and their preparation as a feeder layer for the hESCs (see Support Protocol), the preparation of hESC for plating on to the END2 cells, maintenance of the cocultures during differentiation to cardiomyocytes (see Basic Protocol 1), and the selection and dissociation of the beating areas, as required for further experimentation (see Basic Protocol 2). NOTE: For all procedures described in this unit, tissue culture, reagent preparation and sterilizing facilities are necessary. Class II Biological Hazard Flow Hoods (for END2 passage and mitomycin C treatment) or laminar flow horizontal draft hoods (for hESC passage and transfer to coculture) are used. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 8% CO2 incubator unless otherwise specified. NOTE: Ethical approval and/or institute review may be required for the use of some or all hESC lines. BASIC PROTOCOL 1

SETTING UP hESC AND END2 COCULTURES AND MAINTENANCE OF CULTURE DURING DIFFERENTIATION The method is described for hESCs routinely passaged mechanically by cut-and-paste, but the method is also effective in some, but not all, enzymatically passaged cell lines provided they are not dispersed too rigorously and clumps become very small. The ES cells may then be rather more contaminated with mouse or human feeder cells but these generally fail to attach and are removed at the first medium change.

Materials Culture plates (12-well) with mitomycin C–treated END2 cells (Support Protocol) hESC medium (for hES2 and hES3 cells; see recipe) containing FBS for undifferentiated cell growth hESC medium without FBS, for differentiation Colonies of hESC (UNIT 1C.1) in organ culture dishes Phosphate-buffered saline calcium- and magnesium-free (CMF-PBS; Invitrogen), sterile, 37◦ C Dispase solution, 10 mg/ml in standard hESC medium, freshly prepared and filter sterilized (using a 0.22-µm filter) Antibody against α-actinin (mouse monoclonal; Sigma), optional Antibody against tropomyosin (mouse monoclonal; Sigma), optional 37◦ C, 5% CO2 incubator 35-mm tissue culture dishes Organ culture dishes 1000-µl pipet 200- to 1000-µl pipet tips Stereo dissecting microscope at 4× magnification (with heated stage if possible) Additional reagents and equipment for obtaining culture plates with mitomycin C–treated END2 cells (Support Protocol) and obtaining organ dishes containing colonies of hESC (UNIT 1C.1)

Differentiation of hESCs to Cardiomyocytes

Set up cocultures 1. Refresh mitomycin C–treated END2 cells (see Support Protocol) in 12-well plates with hESC medium without FBS (1 ml per well) at least 1.5 hr before plating the hESC cell pieces.

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2. Prepare dishes for washing undifferentiated hESC colony-pieces by filling (a) six 3.5-cm petri dishes with CMF-PBS (∼5 ml per dish) and (b) two organ culture dishes with 1 ml hESC medium with fetal bovine serum (FBS); place in incubator at 37◦ C and 5 % CO2 . 3. Collect the starting material for two 12-well hESC-END2 coculture plates: twelve organ culture dishes each with 9 to 10 colonies of either hES2 or hES3. Detach hESC colonies from MEFs by adding 0.5 ml dispase solution and placing in the 37◦ C incubator for 7 min. 4. Collect all undifferentiated hESC colonies from the twelve organ culture dishes using a 1000-µl pipet with a 200- to 1000-µl pipet tip. Lift them off the dishes and distribute them for washing over three 35-mm dishes containing CMF-PBS prepared previously. Depending on the number of dishes being used, colonies may stay longer in CMF-PBS (for several minutes). It is advisable to continue with the next step as soon as possible.

5. Transfer the colonies to three new CMF-PBS dishes (to remove MEFs attached to the colonies). 6. Transfer the colonies to two organ culture dishes containing 1 ml standard hESC medium with FBS. 7. Break colonies into pieces by firmly pipetting up and down (one to three times depending on the size of the colonies) against the bottom of the dish using a 1000-µl pipet (at an angle of ∼ 45◦ to 60◦ ) under the stereomicroscope. 8. Transfer small clumps of hESC to two 12-well plates (5 to 10 clumps per well) containing confluent mitomycin C–treated END2 cells (see Support Protocol). This is day 0. At this point, start using the hESC medium without FBS. Cells will attach after 24 hr. Figure 1F.2.1A shows the sizes of the initial clumps and how differentiation proceeds from day 0 to day 12. Clumps of ∼0.3 to 0.5 mm are clearly visible to the naked eye and will attach by day 2 (Fig. 1F.2.1C). The smaller clumps remain in suspension and do not differentiate into cardiomyocytes (Fig. 1F.2.1D).

Differentiate cells 9. Refresh medium on days 5, 8/9, and 11/12 (1 ml per well) after plating, depending on which day coculture was started. Day 8/9 is the Friday before the weekend and day 11/12 is the Monday after the weekend. Refreshment of the medium on day 5 removes all of the smallest aggregates shown in Figure 1F.2.1D.

10. Score beating areas by microscopic examination 12 days after plating. Beating generally starts on days 5 to 7 (Figs. 1F.2.1E,F) and is maximal on day 12. Differentiation is usually accompanied by the formation of cystic bodies (see Figs. 1F.2.1G,H for examples of culture morphology on day 12).

11. Optional. Immunostain with antibodies against α-actinin (mouse monoclonal; dilution 1:800) and/or tropomyosin (mouse monoclonal; dilution 1:400) to count the number of cardiac cells in the beating clumps. This is usually 20% to 25% of the cells. 1. Quantification may alternatively be carried out by immunoblotting (using e.g., troponin I antibodies; rabbit polyclonal AB1627, Chemicon International; dilution 1:100). 2. The efficiency of differentiation towards cardiomyocytes may be variable. Important factors are the initial number of cells per clump (i.e., the size of the clumps; be sure not to make them too small; see Fig. 1F.2.1A for ideal sizes of the clumps; the smallest do

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Differentiation of hESCs to Cardiomyocytes

Figure 1F.2.1

Legend at right.

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not attach and are removed when medium is refreshed) and the FBS batch, even if the differentiation is in serum-free medium. It is important not to refresh the medium too often as the colonies then tend to remain too small. Serum replacement (KO-SR, Invitrogen) contains factors that are inhibitory and so it should not be used. 3. In the authors’ hands, some hESC lines respond best in END2 cocultures [e.g., hES2 and hES3-GFP (Envy)] whilst others respond better when grown as aggregates in END2conditioned medium (e.g., hES3), as described previously for pluripotent P19 embryonal carcinoma cells (van den Eijnden-van Raaij et al., 1991). hESC medium without serum is conditioned for 4 days by a confluent END2 cell monolayer prior to use. NL-HES1 (van de Stolpe et al., 2005) and NL-HES2 (unpub. observ.), both derived on MEFs and NL-HES3 and NL-HES4 (D. Ward, unpub. observ.), derived on human foreskin fibroblast feeders, differentiate both in END2 coculture and as EBs in END2 conditioned medium.

CULTURE OF END2 CELLS AND PREPARATION AS FEEDER CELLS Mouse END2 cells proliferate exponentially in monolayer culture and become contact inhibited at confluence. They have multiple properties of extraembryonic (visceral yolk sac) endoderm, including expression of α-fetoprotein and the cytoskeletal gene ENDOA, which is recognized by the antibody TROMA-1. They promote cardiomyogenesis in P19 embryonal carcinoma cells and mouse ES cells as well as hESC, as described here.

SUPPORT PROTOCOL

NOTE: END2 cells can be obtained from C. Mummery at the Hubrecht Laboratory ([email protected]) following completion of a Material Transfer Agreement on the Web site http://www.niob.knaw.nl.

Materials 0.1% (w/v) gelatin (see recipe) END2 culture medium (see recipe) END2 cells ([email protected]) Phosphate-buffered saline, calcium and magnesium free (CMF-PBS; Invitrogen), sterile Mitomycin C stock (see recipe) Trypsin/EDTA (Invitrogen, no. 25300-054) 25-, 75-, and 175-cm2 tissue culture flasks coated with 0.1% gelatin 37◦ C incubator 12-well plates coated with 0.1% gelatin Cover slips treated with 0.1% gelatin, in a 12-well plate Additional reagents and equipment for counting cells (Phelan, 2006) Figure 1F.2.1 (at left) Morphology of hESC during differentiation to cardiomyocytes. Stereo images of (A) Dispersed H=hES3 colonies immediately after collection and dissociation. Colonies in the correct size range are indicated by arrowheads. (B) Dispersed colonies immediately after plating on END2 cells pretreated with mitomycin C. The END2 cells cover the black background areas but are not visible in stereo images. (C) Cocultures on day 2 showing large colonies that have attached that will go on to start beating; (D) Area of coculture on day 2 with very small colonies/single cells that will not start to beat and will be washed away on day 5 when medium is refreshed; (E) Day 5 of coculture–colonies are beginning to grow; (F) Day 6 of coculture; beating areas may be visible; (G, H) Day 12 of coculture; two views of coculture with many beating areas that will be cut out for dissociation. Arrows indicate beating areas. In general, these areas may be maintained for several days in this state but no new beating areas will appear. After longerterm maintenance in serum-free medium, the culture in general begins to deteriorate and noncardiomyocytes lyse. The authors have maintained isolated beating colonies in serum-containing medium for up to 3 months. (I) Beating embryoid body on day 12 after growth in serum-free END2 conditioned medium. (J) cardiomyocyte stained with α-actinin antibodies after dissociation and re-attachment. Scale bar: 1 mm. Figure 1F.2.1J provided by Stefan Braam.

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Gelatin-coat flasks, plates, or dishes 1. Add 0.1% gelatin to flasks, plates, or dishes at 1 ml/cm2 , and leave for at least 1 hr at room temperature. Just before plating the cells remove gelatin. Routinely passage and culture END2 cells 2. Passage END2 cells twice weekly (Monday and Thursday or Friday; see below). 3. Remove culture medium, rinse twice in CMF-PBS, and add trypsin/EDTA (0.5 ml for 25-cm2 flask; 1.5 ml for 75-cm2 flask; and 4.0 ml for 175-cm2 flask) 4. Leave cells for 5 min at 37◦ C. Check if cells are rounded up/come off the plate and resuspend in END2 culture medium (5 ml for 25-cm2 flask; 10 ml for 75-cm2 ; and 20 ml for 175-cm2 flask). 5. Transfer to new gelatin-coated flasks, using a split ratio of 1:8 for Monday to Friday culture or 1:6 Monday to Thursday, but always check before passage that culture is confluent. In general (if using other days of the week) split 1:8 (seed 8,125 to 10,000 cells/cm2 ) for 4 days, 1:6 (seed 11,000 to 13,000 cells/cm2 ) for 3 days. For T25 flask use ∼7 ml of culture medium, for T75 flask ∼20 ml, and for T175 flask ∼40 ml of culture medium END-2 cells can be transported frozen (dry-ice) or as growing cultures (seeded at low density 1 day before shipment) at room temperature. The culture flask is then completely filled with medium (pH should be adjusted to 7.3) and sealed with Parafilm. The cells will survive for at least a week under these conditions.

Mitomycin C treat END2 cells for hESC coculture 6. On day 1 (preferably Monday), seed a 25-cm2 tissue culture flask coated with 0.1% gelatin with END2 cells in END2 culture medium. END2 cells should be split 1:8 from a confluent flask.

7. On day 5, seed a 175-cm2 flask coated with 0.1% gelatin with END2 cells using all the cells from the previous 25-cm2 flask. 8. Culture to day 8, when the 175-cm2 flask is 100% confluent and ready for mitomycin C treatment. 9. Remove some of the culture medium from the flask, leaving 25 ml. Add 5 µl mitomycin C per ml medium from 2 µg/µl stock solution to the culture medium (final concentration is 10 µg/ml). In this case add 125 µl mitomycin C per flask.

10. Incubate flasks for at least 2.75 hr and maximally 3 hr, at 37 ◦ C. 11. Aspirate medium, wash once with ml END2 culture medium, followed by two washes with CMF-PBS (∼30 to 35 ml of medium/CMF-PBS each time) CAUTION: The waste containing mitomycin C is highly toxic. Handle and dispose with care according to local laboratory guidelines.

12. Trypsinize with 4 ml trypsin/EDTA, and count the cells (Phelan, 2006). This will yield 65,000 to 80,000 cells/cm2 .

Differentiation of hESCs to Cardiomyocytes

13. Plate END2 cells at a density of 175,000 cells/ml in END2 culture medium: 1 ml cell-suspension per well of a 12-well plate precoated with 0.1% gelatin, with or without 15-mm diameter cover slips, as required for further experimentation.

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SELECTION AND DISSOCIATION OF BEATING AREAS Cells in the beating areas of 12-day cocultures can be isolated for further characterization.

BASIC PROTOCOL 2

Materials 12-day cocultures of hES2 or hES3 cells displaying beating areas (Basic Protocol 1) hESC medium with FBS (see recipe) Low-calcium buffer (buffer 1; see recipe) Enzyme buffer (buffer 2; see recipe) KB buffer (buffer 3; see recipe) Scissors 12-well plate 1000-µl pipet 200- to 1000-µl pipet tips Nonpivoting shaker1.5-ml microcentrifuge tube, optional Gelatin-coated (0.1%) coverslips Additional reagents and equipment for obtaining 12-day cocultures of hES2 or hES3 cells displaying beating areas (Basic Protocol 1) 1. Isolate beating areas (three-dimensional structures) from coculture plates (Basic Protocol 1) by cutting them out with scissors and collect excised tissue in standard hESC medium with FBS. 2. To start dissociation, transfer excised tissue pieces to one well of a 12-well plate with 1 ml of low-calcium buffer (buffer 1) using a 1000-µl pipet with a 200- to 1000-µl pipet tip; leave for 30 min at room temperature. IMPORTANT NOTE: After incubation in low-calcium buffer, the three-dimensional structures no longer contract.

3. Transfer three-dimensional structures from low-calcium buffer (buffer 1) into 1 ml enzyme buffer (buffer 2) in a new well of a 12-well plate and incubate for ∼ 45 min at 37 ◦ C for (cover dish with Parafilm before transferring to the CO2 incubator). 4. Transfer cell clumps from enzyme buffer (buffer 2) into 1 ml KB buffer (buffer 3), again in new well of a 12-well plate. Shake gently in KB buffer (buffer 3) for 1 hr at 100 rpm, room temperature, on a nonpivoting shaker. 5. Transfer three-dimensional structures from KB buffer into 1 ml hESC culture medium with 20% FBS to promote attachment and survival. Break up the cell clumps by pipetting up and down (two to four times) against the bottom of the dish or in a sterile 1.5-ml microcentrifuge tube using a 1000-µl pipet. The degree of dissociation required depends on the particular experiments that will be done with the cardiomyocytes: For transplantation into animals or immunofluorescent staining, it is sufficient to obtain a mixture of cell clumps and single cells. For electrophysiology single cells are required. Do not try to get a complete single cell suspension. Even if clumps are still present, there will be many single cells in suspension (see remark below).

6. For electrophysiology and immunofluorescent staining, seed dissociated beating areas in standard hESC medium on gelatin-coated (0.1%) coverslips in culture plates (preferably a 12-well plate) at 37◦ C, as required, between 10,000 and 15,000 cells per well in a 12-well plate. Be careful that the density is not higher than this, otherwise non-cardiomyocytes will overgrow the cardiomyocytes.

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7. Allow cells on coverslips to recover for at least 2 days and up to a week for electrophysiology experiments (Mummery et al., 2003). Change the medium to remove dead cells after 3 to 5 days but be careful not to dislodge attached cells. 8. For transplantation (Mummery et al., 2003) into mouse heart, keep dissociated cells in suspension in hESC medium at 4◦ C on ice. Inject 105 to 106 cells into the left ventricle in a volume of maximally 5 µl of medium. Larger volumes cause scarring of the cardiac tissue. Cardiomyocytes are very fragile! If pipetting is too rigorous, cells will fail to recover either in culture or in vivo.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

END2 culture medium DMEM/F12 (1:1; Invitrogen, no. 31331-028; contains GlutaMAX I) 1× penicillin/streptomycin (from 500× stock solution; Invitrogen, no. 15070-063) 1× MEM Non-Essential Amino Acids (from 100× stock solution; Invitrogen, no. 11140-035) 7.5% (v/v) fetal bovine serum (FBS) Store up to 4 weeks at 4◦ C FBS is always batch tested for optimal growth. Possible suppliers are Cambrex or Greiner. Perbio is the supplier for Hyclone Serum.

Enzyme buffer (buffer 2) 12 ml 1 M NaCl 3 µl 1 M CaCl2 0.54 ml 1 M KCl 0.50 ml 1 M MgSO4 0.50 ml 1 M sodium pyruvate 2 ml 1 M glucose 20 ml 0.1 M taurine 1 mg/ml (final) collagenase A (100 mg total/100 ml buffer) 1 ml 1 M HEPES Adjust to 100 ml using H2 O Adjust pH to 6.9 with NaOH Filter sterilize Divide into 1-ml aliquots Store up to 6 to 12 months at −20◦ C The efficiency of dissociation of the beating clusters into individual cardiomyocytes may depend on the batch of collagenase A used and batch testing may be advisable. The authors use collagenase A from Roche (no. 11088793-001).

Gelatin, 1.0% Dissolve 0.4 g gelatin (Sigma, no. G1890) in 40 ml distilled water in 50-ml tube, cover tube with aluminum foil and autoclave. Use this solution to prepare 0.1% gelatin: dilute 1:10 with distilled water and store the 0.1 % solution up to 6 weeks at 4◦ C (store the 1.0% solution up to 1 year at −20◦ C).

hESC culture medium (for HES2 and HES3 cells) Differentiation of hESCs to Cardiomyocytes

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DMEM (high glucose; Invitrogen, no. 11960-044) containing: 1× L-glutamine (from 100× stock, Invitrogen, no. 25030-024) continued Current Protocols in Stem Cell Biology

1× penicillin/streptomycin (from 200× stock, Invitrogen, no. 15070-063) 1× nonessential amino acids (from 100× stock, Invitrogen, no. 11140-035) 1× insulin, transferrin, selenium (ITS; from 100× stock, Invitrogen, no. 41400045) 1× 2-mercaptoethanol (from 555× stock, Invitrogen, no. 31350-010) 20% fetal bovine serum (FBS; Hyclone/Perbio) Filter medium through Stericup-GV filter unit (Millipore) Store up to 4 weeks at 4◦ C (turnover is usually much faster with regular use) FBS is omitted from the medium for differentiation. FBS is always batch tested for optimal growth. Possible suppliers are Cambrex or Greiner. Perbio is the supplier for Hyclone Serum.

KB buffer (buffer 3) 8.5 ml 1 M KCl 2 mM Na2 ATP 0.50 ml 1 M MgSO4 0.1 ml 1 M EGTA 0.50 ml 1 M sodium pyruvate 5 ml 0.1 M creatine 20 ml 0.1 M taurine 2 ml 1 M glucose, added just prior to use 3 ml 1 M K2 HPO4 (added last) Adjust to 100 ml using H2 O Adjust pH to 7.2 with NaOH Filter sterilize Divide into 1-ml aliquots Store up to 6 to 12 months at −20◦ C When making KB buffer leave out the glucose, otherwise a precipitate forms at −20◦ C; add glucose just prior to use. Add K2 HPO4 as the last step, otherwise a precipitate forms.

Low-calcium buffer (buffer 1) 12 ml 1 M NaCl 0.54 ml 1 M KCl 0.50 ml 1 M MgSO4 0.50 ml 1 M sodium pyruvate 2 ml 1 M glucose 20 ml 0.1 M taurine 1 ml 1 M HEPES Adjust to 100 ml using H2 O Adjust pH to 6.9 with NaOH Filter sterilize Divide into 1-ml aliquots Store up to 6 to 12 months at −20◦ C Mitomycin C stock Prepare mitomycin C stocks by injecting 1 ml of CMF-PBS (using a 3-ml syringe and an 18-G needle) into the mitomycin C stock bottle (Sigma, no. M0503) to dissolve the powder, taking care that all the powder is well dissolved, then transfer the solution (within the syringe) through a sterile 0.22-µm syringe filter into a 1.5-ml cryotube for immediate use or storage in the dark for up to 4 weeks at 4◦ C.

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COMMENTARY Background Information

Differentiation of hESCs to Cardiomyocytes

A continuous supply of human cardiomyocytes would be a valuable resource, not only for screening drugs that specifically target cardiac cells but also for identifying cardiotoxic risk for other, noncardiac drugs. In addition, the production of human cardiac cells bearing specific gene mutations would facilitate the development of improved in vitro methods for testing subsets of disease-related drugs and physiology. In the long term, it is also hoped that cardiomyocyte replacement therapies may become an important part of cardiac medicine as a means of supplementing or restoring contractile force in the failing heart. A prerequisite is the efficient production of large and preferably homogeneous cell populations of cardiomyocytes. Since human embryonic stem cells are pluripotent and have the capacity for indefinite self-renewal, they are considered as a potentially promising source of cardiomyocytes. However, the first report of cardiomyocytes derived from hESC (Kehat et al., 2001) appeared only 3 years after hESC were first derived from blastocyst-stage embryos (Thomson et al., 1998). The differentiation protocols that are effective in inducing hESC to differentiate into cardiomyocytes depend in part on the individual hESC line used and in part on how they are propagated prior to differentiation. Many of the current methods are based on methods developed for mouse ES cells more than two decades ago (Doetschmann et al., 1985). Mouse ES cells start to beat spontaneously in culture if grown for 4 to 10 days as aggregates in suspension. Because these aggregates form an outer layer of (extra) embryonic endoderm they have been termed embryoid bodies (EBs). The endoderm may be important as a source of differentiation signals in EBs since it is known that in normal development, endoderm is essential for and signals to the anterior mesoderm during heart formation. The “spontaneous” conversion of undifferentiated stem cells to cardiomyocytes is generally a low-efficiency process with only 1% to 2% of the cells within a beating EB or cell aggregate being cardiomyocytes. Much of the literature on improving these efficiencies concerns activating specific developmentally relevant signaling pathways whilst the cells are growing as EBs. The wnt and bone morphogenetic protein (BMP) signaling pathways, for example, have proved most potent in the mouse although addition of ascorbic acid, absence of fetal bovine serum, and a variety of

modifications to the growth medium or ways of forming cell aggregates have all been reported to enhance efficiency. Some of these methods have now been tested in hESC but only a few have been effective, possibly indicative of species differences. There are no descriptions to date describing complete conversion of either mouse or human ES cells into cardiomyocytes although genetic selection methods, using cardiac-specific promoters in combination with selection markers, which are proving effective for scale up and selection of cardiomyocytes from mouse ES cells, are presently under development for hESC. Selection and enrichment may contribute usefully to attempts to produce homogeneous human cardiomyocyte populations, but it is generally believed that greatest benefit is still to be gained by finding methods for improving efficiency of differentiation. In the following section, the results that have been obtained using different hESC lines will be described. This may help in facilitating the choice of cell line since, in general, practical reasons (e.g., labor, cost) may limit the number of different hESC lines that any individual laboratory can deal with. Cardiomyocyte differentiation of hESC In the first study to induce cardiomyocyte differentiation of hESC (cell line H9.2), Kehat et al. (2001) used collagenase IV to disperse the cells into small clumps (3 to 20 cells) and grew them for 7 to 10 days in suspension to form EB-like structures but apparently without the distinct outer layer of endoderm cells. After plating these EBs onto gelatin-coated culture dishes, beating areas were first observed in the outgrowths 4 days after plating (i.e.11 to 14 days after the start of the differentiation protocol). A maximum number of beating areas was observed 20 days after plating (i.e., 27 to 30 days of differentiation), with 8.1% of 1884 EBs scored as beating. This spontaneous differentiation into cardiomyocytes in aggregates was also observed by others using different cell lines e.g., H1, H7, H9, H9.1, and H9.2 (Xu et al., 2002). However, in this case ∼70% of the EBs had beating areas after 20 days of differentiation and even by day 8, 25% were beating. A third group also reported spontaneous formation of cardiomyocytes from the H1, H7, H9, and H14 cell lines but here, 10% to 25% of the embryoid bodies were beating after 30 days of differentiation (He et al., 2003). The reasons for these apparent differences in efficiency are not clear. In addition, counting

1F.2.10 Supplement 2

Current Protocols in Stem Cell Biology

beating EBs may not accurately reflect the conversion of hESC to cardiomyocytes since individual EBs may contain significantly different numbers of cardiac cells. Recently, the differentiation of several independent hESC lines BG01, BG02, and HUES-7 has been described (Zeng et al., 2004; Denning et al., 2006). The HUES lines 1 to 17 are of particular interest because they were the first to be derived and maintained by bulk enzymatic passaging using trypsin (Cowan et al., 2004) both on mouse embryonic fibroblast layers or on Matrigel in feeder-free conditions (Denning et al., 2006). Beating cardiomyocytes, immunoreactive for the cardiac marker α-actinin, can be derived by EB formation from these hESC lines. EBs are formed either by mass culture of small clumps of hESC harvested using collagenase IV or by forced aggregation of defined numbers of hESC harvested using trypsin. Both methods are provided as bench protocols here, although compared to EBs formed by the mass culture approach, forced aggregation results in greater EB homogeneity and an up to 13-fold improvement in cardiomyogenesis (Burridge et al., 2007). An alternative method for the derivation of cardiomyocytes from the hES2 (Reubinoff et al., 2000) and hES3 cell lines has been described by Mummery et al. (2002, 2003) and Passier et al. (2005). These cell lines are generally passaged by a “cut-and-paste method” (Reubinoff et al., 2000), although they may also be adapted to enzymatic passage. Beating areas were first observed following coculture of hES2 cells (Mummery et al., 2002, 2003) with a mouse visceral endoderm-like cell-line (END2). Endoderm plays an important role in the differentiation of cardiogenic precursor cells that are present in the adjacent mesoderm in vivo. Earlier coculture of END2 cells with mouse P19 embryonal carcinoma (EC) cells, a mouse embryonal carcinoma cell line with pluripotent differentiation properties, and with mESC had already shown that beating areas appeared in aggregated cells and that culture medium conditioned by the END2 cells contained cardiomyogenic activity (van den Eijnden-van Raaij et al., 1991; Mummery et al., 2002). For the derivation of cardiomyocytes from hESC, mitotically inactivated END2 cells were seeded on a 12-well plate and cocultured with the hESC line HES-2 (Fig. 1F.2.1). This resulted in beating areas in ∼35% of the wells after 12 days in coculture. Whilst these methods appear to be effective, all produce cardiomyocytes at low efficiency. Several potential cardiogenic fac-

tors described as effective in mouse ES cells, have been tested in hESC. No significant improvement in cardiomyocyte differentiation has been achieved by adding DMSO, retinoic acid (Kehat et al. 2001; Xu et al., 2002), or BMP-2 (Mummery et al., 2003; Pera et al., 2004). It is not clear whether these factors do not play a role in cardiac differentiation of hESC, or whether differentiation protocols were not optimal. One factor that has been described as enhancing cardiomyocyte differentiation of hESC is the demethylating agent 5 deoxyazacytidine. Treatment of hESC aggregates with 5 deoxyazacytidine (a concentration of 1 and 10 µM 5-deoxyazacytidine was used by Xu et al., 2002) induced enhanced cardiomyocyte differentiation and upregulated the expression of cardiac α-myosin heavy chain, as determined by real time RTPCR, up to two-fold (Xu et al., 2002) although this does not routinely work for all hESC lines. The presence of fetal bovine serum during differentiation also has important effects on differentiation efficiency. In most reports to date, serum has been present in the culture medium, most probably because absence of serum is reported to be detrimental to maintenance of primary cardiomyocytes (Piper et al., 1988). However, serum may contain inhibitory factors and differentiation efficiency has been described as being dependent on serum batch. For example, Sachinidis et al. (2003) observed a 4.5-fold upregulation in the percentage of beating mouse EBs after changing to a serum-free differentiation medium. The authors also recently observed a greater than 20fold increase in cardiomyocyte yield in hES2 + END2 cocultures in serum-free medium (Passier et al., 2005) which was further enhanced by ascorbic acid. The phenotype of the majority (∼90%) of cardiomyocytes derived using this coculture protocol show greatest similarity to human fetal ventricular cells, although atrial and pacemaker-like cells are also observed (Mummery et al., 2003). This serum-free protocol is described here for both hES2 and hES3 cells. The method has also proved effective on all four recently derived hESC lines (see van de Stolpe et al., 2005 for NL-HES1) but was ineffective in BG01 (Burridge et al., 2007). hES3 cells also differentiate to cardiomyocytes as aggregates in suspension in the presence of serum-free medium conditioned by END2 cells for 4 days. Here the method used for generating and characterizing cardiomyocytes from hES2 and hES3 has been described, as well as a method for dissociation of cardiomyocytes into

Mesodermal Lineages

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single-cell suspensions, which is useful both for characterizing cells by antibody staining (FACS analysis) and electrophysiological measurements. The dissociated cells are also suitable for transplantation into (animal) hearts.

Critical Parameters It has proven essential for the authors that the hESC are induced to differentiate in small clumps or undergo some sort of forced aggregation step (this is particularly important for cell lines that are passaged enzymatically, rather than mechanically). Burridge et al. (2007) have also recently emphasized the difference in response of different cell lines in this respect. For the hESC + END2 coculture protocol described and for the growth of aggregates in END2-conditoned medium, it is essential that the aggregates are neither too small nor too large. Practice will give a feeling for the best method of making the clumps. The authors have derived four hESC lines under different conditions, two on mouse embryonic feeder cells in standard hESC medium, as given, and two on human foreskin fibroblasts in KO-DMEM and serum replacement (SR). All respond by forming cardiomyocytes in both coculture and suspension methods but the efficiency improved after passage 10. Serum or serum replacement inhibits differentiation when present during the first week (Passier et al., 2005) but can be added thereafter to enhance longer-term survival. Ascorbic acid also enhances differentiation efficiency in hES2 cells (Passier et al., 2005) but several other growth factors that have been implicated in mesoderm or cardiomyocyte differentiation (BMP2 or 4, FGF, PDGF, and oxytocin) were without effect.

Troubleshooting

Differentiation of hESCs to Cardiomyocytes

The most frequent problems encountered are that individual hESC lines may respond variably from week-to-week or that particular hESC lines will not form cardiomyocytes at all, or only very inefficiently. By following protocols very precisely and changing reagents as little as possible (FBS and SR are batch tested and bought in at least a year or more supply at once) the authors have been able to produce cardiomyocytes very reproducibly on a weekly basis for several years. Developing routine in the methodology is also of importance since it can be a source of variability between operators. Some cell lines may not appear to form beating cardiomyocytes at all. It may still be worth staining for contractile

proteins since very immature cells that have not yet started to beat may have formed. They can sometimes be activated by tapping the dish gently or electrical pacing with a microelectrode. In collaboration with others, the authors have tried the END2 coculture protocol on BG01 without success and on H9 cells with only low efficiency of differentiation. If at all possible, obtaining a hESC line with proven ability to form cardiomyocytes is a very useful positive control for methodology and operator. hES2 and hES3 are both on the NIH-approved list and may be used with U.S. and EU funding.

Anticipated Results For electrophysiological studies relatively few, single cardiomyocytes are needed, and they can be produced fairly easily from several hESC lines. However, for other types of study (e.g., proteomics, microarray, and transplantation to the hearts of animals) much larger numbers of cardiomyocytes are required and the cultures are preferably substantially enriched. Using the coculture method on hES3 or hES3-GFP (Envy; Costa et al., 2005) the authors can produce up to 5 × 106 differentiated hESCs dissociated from beating aggregates of which 20% to 25% are cardiomyocytes on a weekly basis (depending on the efficiency of differentiation, six to twelve 12-well plates of hESC-END2 cocultures are required). Selection of beating aggregates results in complete separation from the END2 cells. Aggregates thus produced have been used successfully for microarray analysis (Beqqali et al., 2006), despite the presence of other cell types, and have shown a pattern of gene expression changes that would be expected from the sequential differentiation of hESC first to nascent mesoderm, then cardiac progenitors, and finally differentiated cardiomyocytes with a fetal phenotype. The arrays also allowed the identification of novel genes associated with cardiomyogenesis in human cells. Production can be scaled up by culturing the cells as aggregates/EBs in END2-conditioned medium although our analysis of the cardiomyocyte characteristics under these conditions has not been as extensive as for cardiomyocytes resulting from coculture.

Time Considerations The basic maintenance of hESC is a fairly labor-intensive process and the authors have a group of three technicians running a small core facility for the production of undifferentiated hESC and cardiomyocytes (or cardiomyocyte cell suspensions). In practice, two independent

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hESC cultures are run in parallel, one passaged on Wednesdays and one on Thursdays. MEFs between passage 4 to 6 (depending on batch) are generally cultured from Thursday to Monday or Thursday to Tuesday, in both cases with an extra passage on Friday. They are treated with mitomycin C on Mondays and Tuesdays for plating hESC on Wednesday and Thursdays (as above). Medium on hESC cultures is refreshed every day (takes about 1 hr but depends on how many cultures there are) except Saturday. END-2 cells between passage 10 and about 30 to 35 are cultured in flasks and treated with mitomycin C on Mondays to prepare the plates for cocultures. The cocultures can be started on either Wednesday and/or Thursday, as part of standard passage, and the coculture refreshment is on day 5, 8/9 and day 11/12 (depending on whether coculture was initiated on Wednesday or Thursday, refreshment takes place on the Friday before or the Monday after the weekend in the second week of differentiation). On day 12 (i.e., Monday or Tuesday, depending when the coculture was initiated), the development of beating aggregates is maximal and aggregates are collected for analysis or dissociation. The most labor-intensive steps are collection and dissociation of cells/beating areas on Mondays and Tuesdays, on day 13 of differentiation (one person the whole morning on each day) and the passage of hESC to MEFs or END2 coculture (one person the whole day, depending on how many cells are required for the next set of experiments). This regime allows reliable production of cardiomyocytes in general although (for unexplainable reasons) cardiomyocyte yields may be substantially higher or lower than expected.

Cowan, C.A., Klimanskaya, I., McMahon, J., Atienza, J., Witmyer, J., Zucker, J.P., Wang, S., Morton, C.C., McMahon, A.P., Powers, D., and Melton, D.A. 2004. Derivation of embryonic stem-cell lines from human blastocysts. N. Engl. J. Med. 350:1353-1356.

Literature Cited

Passier, R., Ward-van Oostwaard, D., Snapper, J., Kloots, J., Hassink, R., Kuijk, E., Roelen, B., Brutel de la Riviere, A., and Mummery, C. 2005. Increased cardiomyocyte differentiation from human embryonic stem cells in serum-free cultures. Stem Cells 23:772-780.

Beqqali, A., Kloots, J., Ward-van Oostwaard, D., Mummery, C., and Passier, R. 2006. Genomewide transcriptional profiling of human embryonic stem cells differentiating into cardiomyocytes. Stem Cells 24:1956-1967. Burridge, P.W., Anderson, D., Priddle, H., Barbadillo Munoz, M.D., Chamberlain, S., Allegrucci, C., Young, L.E., and Denning, C. 2007. Improved human embryonic stem cell embryoid body homogeneity and cardiomyocyte differentiation from a novel V-96 plate aggregation system highlights interline variability. Stem Cells 25:929-938. Costa, M., Dottori, M., Ng, E., Hawes, S.M., Sourris, K., Jamshidi, P., Pera, M.F., Elefanty, A.G., and Stanley, E.G. 2005. The hESC line Envy expresses high levels of GFP in all differentiated progeny. Nat. Methods 2:259-260.

Denning, C., Allegrucci, C., Priddle, H., BarbadilloMunoz, M.D., Anderson, D., Self, T., Smith, N.M., Parkin, C.T., and Young, L.E. 2006. Common culture conditions for maintenance and cardiomyocyte differentiation of the human embryonic stem cell lines, BG01 and HUES-7. Int. J. Dev. Biol. 50:27-37. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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. He, J.Q., Ma, Y., Lee, Y., Thomson, J.A., and Kamp, T.J. 2003. Human embryonic stem cells develop into multiple types of cardiac myocytes: Action potential characterization. Circ. Res. 93:3239. Kehat, I., Kenyagin-Karsenti, D., Snir, M., Segev, H., Amit, M., Gepstein, A., Livne, E., Binah, O., Itskovitch-Eldor, J., and Gepstein, L. 2001. Human embryonic stem cells can differentiate into myocytes with structural and functional properties of cardiomyocytes. J. Clin. Invest. 108:407441 Mummery, C., Ward, D., van den Brink, C.E., Bird, S.D., Doevendans, P.A., Opthof, T., Brutel de la Riviere, A., Tertoolen, L., van der Heyden, M., and Pera, M. 2002. Cardiomyocyte differentiation of mouse and human embryonic stem cells. J. Anat. 200:233-242. Mummery, C., Ward-van Oostwaard, D., Doevendans, P., Spijker, R., van den Brink, S., Hassink, R., van der Heyden, M., Opthof, T., Pera, M., de la Riviere, A.B., Passier, R., and Tertoolen, L. 2003. Differentiation of human embryonic stem cells to cardiomyocytes: Role of coculture with visceral endoderm-like cells Circulation 107:2733-2740.

Pera, M.F., Andrade, J., Houssami, S., Reubinoff, B., Trounson, A., Stanley, E.G., Ward-van Oostwaard, D., and Mummery, C. 2004. Regulation of human embryonic stem cell differentiation by BMP-2 and its antagonist noggin. J. Cell Sci. 117:1269-1280. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A3F.1-A.3F.18. Piper, H.M., Jacobson, S.L., and Schwartz, P. 1988. Determinants of cardiomyocyte development in long-term primary culture. J. Mol. Cell Cardiol. 20:825-835.

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Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotech. 18:399-404. Sachinidis, A., Gissel, C., Nierhoff, D., HipplerAltenburg, R., Sauer, H., Wartenberg, M., and Hesheler, J. 2003. Identification of plateletderived growth factor BB as cardiogenesisinducing factor in mouse embryonic stem cells under serum free conditions Cell Physiol. Biochem. 13:423-429. Thomson, J.A., Itskovitch-Eldor, J., Shapiro, S.S., Waknitz, M.A., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. van den Eijnden-vanRaaij, A.J., van Achterberg, T.A., van der Kruijssen, C.M., Piersma, A.H., Huylebroeck, D., de Laat, S.W., and Mummery, C.L. 1991. Differentiation of aggregated murine P19 embryonal carcinoma cells is induced by a novel visceral-endoderm specific FGF-like factor and inhibited by activin A. Mech. Dev. 33:157-165. van de Stolpe, A., van den Brink, S., van Rooijen, M., Ward-van Oostwaard, D., van Inzen, W.,

Slaper-Cortenbach, I., Fauser, B., van den Hout, N., Weima, S., Passier, R., Smith, N., Denning, C., and Mummery, C.L. 2005. Human embryonic stem cells: Towards therapies for cardiac disease. Derivation of a Dutch human embryonic stem cell line. Reprod. Biomed. Online 11:476486. van Laake, L.W., Passier, R., Monshouwer-Kloots, J., Humbel, B.M., Lips, D.J., Freund, C., den Ouden, K., Ward-van Oostwaard, D., Korving, J., Tertoolen, L.G., van Echteld, C.J., Doevendans, P.A., and Mummery, C.L. 2008. Human embryonic stem cell-derived cardiomyocytes survive and mature in the mouse heart and transiently improve function after myocardial infarction. Stem Cell Research (in press). Xu, C., Police, S., Rao, N., and Carpenter, M.K. 2002. Characterization and enrichment of cardiomyocytes derived from human embryonic stem cells. Circ. Res. 91:501-508. Zeng, X., Miura, T., Luo, Y., Bhattacharya, B., Condie, B., Chen, J., Ginis, I., Lyons, I., Mejido, J., Puri, R.K., Rao, M.S., and Freed, W.J. 2004. Properties of pluripotent human embryonic stem cells BG01 and BG02. Stem Cells 22:292-312.

Differentiation of hESCs to Cardiomyocytes

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Current Protocols in Stem Cell Biology

Isolation of Hematopoietic Stem Cells from Mouse Embryonic Stem Cells

UNIT 1F.3

Shannon L. McKinney-Freeman,1, 2 Olaia Naveiras,1, 2 and George Q. Daley1, 2, 3 1

Division of Pediatric Hematology/Oncology, Children’s Hospital, Boston, Massachusetts Department of Biochemistry and Molecular Pharmacology, Harvard Medical School, Boston, Massachusetts 3 Harvard Stem Cell Institute, Cambridge, Massachusetts 2

ABSTRACT This unit describes a protocol for the isolation of cells from murine embryonic stem cells with hematopoietic stem cell activity, defined by the ability to reconstitute, long term, multiple lineages of the hematopoietic system of lethally irradiated mice. The protocol subjects hematopoietic progenitors specified in differentiating embryoid bodies to ectopic HoxB4 expression (delivered via retroviral infection), followed by coculture and expansion on OP9 stromal cells in the presence of hematopoietic cytokines for 10 days. The protocol results in the generation of hundreds of millions of cells that can rescue mice from lethal irradiation. Although little is known about the phenotype and frequency of the actual hematopoietic stem cell–like cell within the population of cells generated by this protocol, the protocol establishes a system in which these cells can be further studied and the results ultimately translated to the human system. Curr. Protoc. C 2008 by John Wiley & Sons, Inc. Stem Cell Biol. 4:1F.3.1-1F.3.10.
Keywords: embryonic stem cell r mESC r hematopoietic stem cell r HSC r blood r bone marrow transplantation r HoxB4

INTRODUCTION This unit describes a protocol for the derivation from murine embryonic stem cells (mESC) of cells with hematopoietic stem cell (HSC) activity. HSC are functionally defined by their ability to maintain the hematopoietic compartment of an organism for its entire life. The cells isolated via the following protocols are capable of restoring the entire hematopoietic compartment of mice that have been subjected to lethal irradiation and, thus, meet the functional definition of HSC. The following method for the derivation of ESC-HSC is divided into four protocols: the differentiation of ESC into embryoid bodies (EBs; Basic Protocol 1), infection of EBderived cells with retroviral HoxB4 (Basic Protocol 2), expansion of infected EB-derived cells on OP9 stromal cells (Basic Protocol 3), and finally, hematopoietic reconstitution of irradiated mice (Basic Protocol 4). A Support Protocol addresses the culture of OP9 cells that are used as feeder cells for infection and growth of infected EB-derived cells. NOTE: The following procedures are all performed in a laminar-flow hood. NOTE: Aseptic technique must be followed for all of the following procedures, and all solutions and reagents should be sterile. NOTE: All cell incubations are performed in a humidified chamber at 37◦ C and 5% CO2 . NOTE: All cell counts refer to viable cells as defined by trypan blue exclusion. NOTE: Unless otherwise indicated, all centrifugations are 5 min at 500 × g, room temperature.

Embryonic and Extraembryonic Stem Cells

Current Protocols in Stem Cell Biology 1F.3.1-1F.3.10 Published online February 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f03s4 C 2008 John Wiley & Sons, Inc. Copyright


1F.3.1 Supplement 4

BASIC PROTOCOL 1

DIFFERENTIATION OF EMBRYONIC STEM CELLS AS EMBRYOID BODIES It is well documented that cell derivatives of the three major germ lineages develop when ESC are induced to differentiate as embryoid bodies (EBs) in culture (Keller, 2005). EBs are three-dimensional cavitating structures in which blood differentiation can be grossly visualized by the formation of hemoglobinized “blood islands” after more than 6 days of differentiation. In this phase of the protocol, ESC are differentiated as EBs for 6 days, using a commercially available serum prescreened for inducing the specification of hematopoietic progenitors in differentiating ESC, as previously described (Kyba et al., 2002; Wang et al., 2005).

Materials mESC growing as a confluent culture on an MEF feeder layer (see UNIT 1C.3) in a 25-cm2 flask Phosphate-buffered saline (PBS, Invitrogen, cat. no. 20012-027) 0.25% (w/v) trypsin (Invitrogen, cat. no. 15090-046) Differentiation medium (see recipe) 0.4% (w/v) trypan blue (Sigma cat. no. T8154) 25-cm2 tissue culture flask 15- and 50-ml conical tubes Hemacytometer Inverted tissue culture microscope Multichannel micropipettor (e.g., ePet by BioHit, VWR), capable of accurately dispensing 15 µl 10- and 15-cm nonadherent petri dishes Plate shaker (e.g., orbital shaker by Ikaworks, VWR) Harvest mESC 1. Harvest a confluent flask of murine ESCs by washing once with 5 ml PBS followed by the application of 0.5 ml of 0.25% (w/v) trypsin for 1 to 2 min at 37◦ C. 2. Add 5 ml differentiation medium and triturate with a 5-ml pipet three to five times to dissociate ESC colonies. 3. Transfer the cells to a 25-cm2 flask and incubate 45 min. This step depletes the majority of feeder cells.

4. Harvest the supernatant and collect ESC by centrifugation. 5. Count viable cells in an aliquot, using an inverted microscope, hemacytometer, and 0.4% trypan blue (see UNIT 1C.3). 6. Resuspend the feeder cell–depleted ESC at 333,333 cells/50 ml differentiation medium (100 ESC/15 µl).

Plate mESC to form EBs 7. Using a multichannel micropipettor, densely plate 15-µl drops of cell suspension onto the bottoms of two to five 15-cm nonadherent petri dishes (see Fig. 1F.3.1A,B). 8. Gently flip the plates to invert the drops (Fig. 1F.3.1C,D). Incubate 48 hr. 9. Collect day 2 EBs (Fig. 1F.3.1E) by gently swirling the plates and transferring the medium with a 5-ml pipet from up to five plates to a single 50-ml conical tube. Rinse the plates with 4 to 6 ml PBS and add to same tube. Isolation of HSC from mESC

DO NOT pool more than five 15-cm2 petri dishes/50-ml tube. The EBs will grow as they continue to differentiate, and if they are too dense, they may form clumps.

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Current Protocols in Stem Cell Biology

Figure 1F.3.1 Photographs of embryoid body (EB) preparation and OP9 expansion. (A) Using a multichannel pipettor, pipet 15-µl drops of ESC suspended in differentiation medium in rows until the plate is full. (B) Representative full 15-cm petri dishes. (C) Gently flip plates to invert drops. (D) Representative inverted plates. Plates can be stacked during incubation. (E) Day 2 EB. (F) Day 6 EBs. (G) Typical day 7 OP9 colony. Note that the colonies are a heterogeneous mixture of large and small cells. (H) Day 10 OP9 expansion. At this point, the culture becomes a confluent bed of large and small cells with many cells also floating in the medium. All photomicrographs are 100× magnification.

10. Allow the pooled day 2 EBs to settle by gravity (∼10 min). 11. Aspirate the supernatant without disturbing the EBs. Gently resuspend in 10 ml differentiation medium and transfer to 10-cm nonadherent petri dish. 12. Incubate the 10-cm petri dish 2 days on a shaker set to 50 rpm. 13. On day 4 of differentiation, swirl the plate to concentrate the EBs in the center of the dish. Carefully remove 50% (5 ml) of the medium without disturbing the day 4 EBs. 14. Add 5 ml fresh differentiation medium and return the plate to the incubator (with shaking) for 2 more days.

Embryonic and Extraembryonic Stem Cells

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BASIC PROTOCOL 2

INFECTION OF DAY 6 EMBRYOID BODY CELLS WITH MSCV-HoxB4-IRES-GFP Ectopic HoxB4 expression is well documented to induce the expansion of hematopoietic progenitors in both bone marrow and differentiating ESC in a nontransforming manner (Sauvageau et al., 1995). In this phase of the protocol, ectopic HoxB4 is expressed in day 6 EB-derived cells by retroviral infection with MSCV-HoxB4-IRES-GFP. This vector allows one to track ESC-derived hematopoietic engraftment in recipient mice using GFP.

Materials Day 6 EBs (Basic Protocol 1) Phosphate-buffered saline (PBS; Invitrogen, cat. no. 20012-027) Dissociation enzymes (see recipe) Enzyme-free dissociation buffer (Invitrogen, cat. no. 13151-014) 10% IMDM (see recipe) 0.4% (w/v) trypan blue (Sigma cat. no. T8154) OP9 stromal cells: plated in 6-well tissue-culture plates 24 hr prior to use (Support Protocol) MSCV-HoxB4-IRES-GFP viral supernatant: ecotrophic pseudotype prepared according to standard techniques (Pear, 1996) and titered (Cepko, 1996) Protamine sulfate (Sigma, cat. no. P3369) 10% IMDM + cytokine cocktail (see recipe) 15-ml conical tubes Water bath, 37◦ C Inverted tissue culture microscope Hemacytometer Centrifuge with rotor adapted for 6-well plates Dissociate EBs to single cells 1. Transfer day 6 EBs (Fig. 1F.3.1F) to 15-ml conical tubes and allow them to settle by gravity. 2. Aspirate the medium and resuspend in 10 ml PBS. Allow the EBs to resettle by gravity. 3. Aspirate the PBS. 4. Add 250 µl dissociation enzymes and 1 ml PBS. Incubate 20 min in a 37◦ C water bath, occasionally swirling the tube. 5. Add 8 ml enzyme-free dissociation buffer. Triturate with a 5-ml pipet until EBs are fully dissociated. Collect the cells by centrifugation. The mixture will become cloudy with cells as EBs dissociate. Placing the pipet tip against the bottom of the conical tube during trituration will create a shear force that greatly facilitates dissociation. Do not overtriturate. Approximately 10 cycles up and down the pipet should be enough to create a single-cell suspension.

6. Resuspend the cells in 5 ml of 10% IMDM and count viable cells in an aliquot using an inverted microscope, 0.4% trypan blue, and a hemacytometer (see UNIT 1C.3). Between day 4 and day 6 of differentiation, EB cavitation results in a large amount of apoptosis and cell death. Thus, at day 6, upwards of 20% of EB-derived cells may stain with trypan blue.

Isolation of HSC from mESC

Infect day 6 EB-derived cells with MSCV-HoxB4-IRES-GFP 7. For retroviral infection, plate 100,000 cells/well in four 6-well plates preplated with OP9 stroma cells the previous day.

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8. Add an appropriate volume of viral supernatant and 10% IMDM to achieve a multiplicity of infection (MOI) of 5 to 10/well. 9. Add protamine sulfate to a final concentration of 8 µg/ml. Polybrene could also be used, but protamine sulfate is less toxic and works well.

10. Centrifuge the plates 90 min at 1400 × g, room temperature. It is common to employ a “spin infection” for infecting cells with retrovirus (see Pear, 1996).

11. Incubate 4 to 6 hr at 37◦ C. 12. Using a pipettor, harvest the supernatant from the plates into 50-ml conical tubes and collect the cells by centrifugation. 13. Resuspend the cell pellet (may be small) in 48 ml of 10% IMDM + cytokine cocktail. Dispense 2 ml/well of the resuspension to the four 6-well plates in which infection was performed. Always prepare 10% IMDM + cytokine cocktail fresh at the time of use. The authors have experimentally determined that the best results are obtained when the cells spend minimal time in the viral supernatant and are placed quickly into medium with cytokines. Since retrovirus has a short half-life, 4 to 6 hr in the viral supernatant is more than sufficient for a good infection and expansion. This replating step works well.

14. Return the plates to the incubator and proceed to Basic Protocol 3.

OP9 STROMAL CELL CULTURE OP9 cells are used as feeder cells for infection and growth of infected EB-derived cells. The OP9 primary stromal cell line is derived from the calvaria (skullcap) of M-CSF deficient mice and the line is well documented to support the hematopoietic differentiation of mESC (Nakano et al., 1994).

SUPPORT PROTOCOL

Materials OP9 stromal cells (ATCC #CRL-2749) 20% α-MEM (see recipe) 6-well tissue-culture plates 1. Maintain OP9 stromal cells according to standard protocols (see Nakano et al., 1994) in 20% α-MEM. 2. Split at no more than 1:3 at 80% confluence (∼4 to 5 days). OP9 cells will change properties when grown to confluence.

3. At 24 hr prior to infection of day 6 EB-derived cells, plate 25,000 OP9 cells/well of four 6-well plates in 2 ml of 20% α-MEM per well.

EXPANSION OF INFECTED EB-DERIVED CELLS ON OP9 STROMA In this next phase of the isolation of hematopoietic stem cells, infected day 6 EB-derived cells are allowed to expand on OP9 stroma in the presence of a cytokine cocktail, prior to transplantation into recipient animals.

BASIC PROTOCOL 3

Materials Infected mESC cultures grown on OP9 stromal cells in 6-well plates (Basic Protocol 2) 0.05% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25300-062) 10% IMDM + cytokine cocktail (see recipe) Current Protocols in Stem Cell Biology

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50-ml conical tubes 75-cm2 tissue culture flask 1. Incubate the mESC plates for 7 days postinfection. Colonies should be apparent by day 4 postinfection and large and well formed by day 7 (Fig. 1F.3.1G). A robust expansion should yield 25 to 50 colonies/well on day 7.

2. On day 7 add a few drops (∼200 µl) of 0.5% trypsin/EDTA to each well, incubate 2 to 3 min, and collect all of the cells (infected ESCs and OP9 cells) into a 50-ml conical tube. Wash the wells with 1 to 2 ml PBS (if necessary), add the wash to the collection tube, and centrifuge. DO NOT discard the supernatant or PBS washes employed during trypsin treatment. At this point in the culture, some colonies may be only loosely adherent, and there are many cells floating in suspension. Collect and pool all washes and supernatant to ensure that no potentially valuable cells are discarded.

3. Pool and resuspend the cells in 8 ml fresh 10% IMDM + cytokine cocktail. Distribute among four 75-cm2 flasks and add an additional 13 ml of 10% IMDM + cytokine cocktail to each flask. Incubate for 3 additional days. 4. On day 10 (Fig. 1F.3.1H), collect and pool cells from all 75-cm2 flasks via treatment with trypsin followed by centrifugation (see step 2). More than 90% of day 10 expanded cells are GFP+ (as determined by FACS or fluorescence microscopy), indicating that they derive from the MSCV-HoxB4-IRES-GFP infected cells. DO NOT discard the supernatant or PBS washes employed during trypsin treatment. At this point in the culture, some colonies may be only loosely adherent and there are many cells floating in suspension. Collect and pool all washes and supernatant to ensure that no potentially valuable cells are discarded. A good expansion should yield between 40–50 × 106 cells/original 6-well plate infected, for a total of 160–200 × 106 cells. BASIC PROTOCOL 4

RECONSTITUTION OF RECIPIENT MICE HSC are defined functionally by their ability to maintain and replenish the hematopoietic system. Thus, the most rigorous assay for HSC functionality is the rescue of an animal model from lethal irradiation and the restoration of its hematopoietic compartment. In the final phase of the protocol, the OP9-expanded cell population is transplanted into lethally irradiated mice. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

Materials OP9-expanded cells (Basic Protocol 3) Serum-free IMDM (Invitrogen) or PBS (Invitrogen) Recipient mice: 6- to 8-week-old Rag-2−/− γc−/− mice weighing 15 to 22 grams (Taconic Farms) Sterile H2 O

Isolation of HSC from mESC

Cesium-source γ-irradiator 1-ml syringe 301/2 -G needles Autoclaved mouse cages

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Additional reagents and equipment for evaluating cells and organs by fluorescence microscopy or flow cytometry (e.g., see Robinson et al., 2008; http://www.cyto. purdue.edu) 1. Resuspend the OP9-expanded cells in serum-free IMDM or PBS (106 cells/300 µl for injection). 2. Ablate the hematopoietic compartment of Rag-2−/− γc−/− mice (or other suitable murine recipient) weighing 15 to 22 grams by exposure to 9.25 gy of irradiation applied in two doses separated by 2.5 hr. It is CRITICAL that the weight of the animals fall between 15 and 22 grams at time of irradiation. If they are larger than 22 grams, 9.25 gy will not be a sufficient dose of irradiation to ensure the appropriate ablation, and cells may not mediate rescue from lethal irradiation and/or engraft the hematopoietic compartment. If transplanting larger animals, the appropriate irradiation dose MUST be determined experimentally.

3. Deliver 5–10 × 106 cells/animal via tail vein injection using a 1-ml syringe with 301/2 -G needles. A minimum dose of 5–10 × 106 cells/animal is required to guarantee rescue from lethal irradiation. If fractionating cells, inject 5–10 × 106 cell equivalents (e.g., if the population of interest represents 10% of unfractionated pool of cells, inject 5–10 × 105 cells/animal). If injecting unfractionated cells, ALWAYS inject via tail vein to avoid teratoma formation in the eye orbit, which may result if high numbers of unfractionated cells are delivered retro-orbitally.

4. Maintain Rag-2−/− γc−/− recipients in autoclaved cages on sterile water. Depending on the cleanliness of the animal facility, posttransplant animals may require administration of acidified water or trimethiprim-sulfasoxazole (Septra) for the 5 weeks following lethal radiation.

5. Assess reconstitution of the hematopoietic compartment of recipients by examining the peripheral blood and hematopoietic organs for the presence of GFP+ cells by fluorescent microscopy or flow cytometry. The most ideal experiments exploit the CD45 congenic system to discriminate OP9expanded cells from radio-resistant host: these cells can be transplanted into recipients expressing the opposite CD45 allelic variant. As GFP expression tends to be silenced by one year posttransplant, the CD45 approach is critical for observing the repopulation of ESC-derived cells at very late time points posttransplant. The CD45 congenic system is classically exploited in the hematopoietic stem cell world because all hematopoietic cells express CD45, and it is convenient. However, one could use any congenic system that would allow one to differentiate donor from host (e.g., Ly-9, Thy-1, MHC).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

α-MEM, 20% α-MEM (Invitrogen) containing: 20% (v/v) heat-inactivated fetal bovine serum (Invitrogen, cat. no. 16000-044) 1% (v/v) 100× penicillin/streptomycin/glutamine (Invitrogen, cat. no.10378-016) Store up to 1 month at 4◦ C

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Differentiation medium IMDM (Invitrogen) containing: 200 µg/ml holo-transferrin (Sigma, cat. no. T1283-1G) 4.5 mM monothioglycerol (Sigma, cat. no. M2172) 1% (v/v) 100× penicillin/streptomycin/glutamine (Invitrogen, cat. no. 10378-016) 15% (v/v) differentiation serum (StemCell Technologies, cat. no. 06952) 50 µg/ml ascorbic acid, cell culture tested (Sigma, cat. no. 4403) Store up to 1 month at 4◦ C Dissociation enzymes Resuspend the following in 50 ml DMEM (Invitrogen): 500 mg collagenase IV (Invitrogen, cat. no. 17104-019) 1 g hyaluronidase (Sigma, cat. no. H2126) 40,000 U DNAse (Sigma, cat. no. D4527) Store at −20◦ C in 500-µl aliquots Avoid repeated freeze/thaws IMDM, 10% IMDM (Invitrogen) containing: 10% (v/v) heat-inactivated fetal bovine serum (Invitrogen, cat. no. 16000-044) 1% (v/v) 100× penicillin/streptomycin/glutamine (Invitrogen, cat. no.10378-016) Store up to one month at 4◦ C IMDM, 10% + cytokine cocktail Prepare the necessary volume of 10% IMDM (see recipe) and supplement with: 100 ng/ml recombinant human Flt3L (Peprotech, cat. no. 300-19) 100 ng/ml recombinant human SCF (Peprotech, cat. no. 300-07) 40 ng/ml recombinant human thrombopoietin (hTPO; Peprotech, cat. no. 300-18) 40 ng/ml recombinant murine VEGF (Peprotech cat. no. 450-32) 10% IMDM + cytokine cocktail must be made fresh for every experiment. However, the cytokines can be stored indefinitely as 100× stock solutions in 500-µl aliquots at −20◦ C.

COMMENTARY Background Information

Isolation of HSC from mESC

HSC transplantation is arguably the most successful and widely applied cell-based therapy used in medicine. However, many patients who might benefit from this therapy lack access to suitable donor material. Thus, coaxing ESC to produce HSC has been attempted many times since these cells were first generated over 20 years ago. Attempts to generate HSC directly from EBs has largely resulted in short-lived hematopoietic repopulation, failure to robustly restore all hematopoietic lineages, or a lack of follow-up studies to verify the robustness of the reported protocol (Brent et al., 1990; Chen et al., 1992; Muller and Dzierzak, 1993; Palacios et al., 1995). Most recently, tantalizing progress has been achieved on the derivation of cells with in vivo HSC activity directly from EBs; c-kit+ CD45+ EBderived cells were able to mediate high levels of multilineage, long-term reconstitution

in allogenic recipients without obvious rejection problems (Burt et al., 2004). However, in a follow-up study, only modest reconstitution was achieved via the application of this protocol, and the highest levels of reconstitution were apparent only when cells were delivered directly into the bone marrow cavity of femurs (Verda et al., 2007). Intrafemoral injection is a laborious and technically challenging technique. These data also suggest that EB-derived repopulating cells have a bone marrow homing deficiency, perhaps a barrier to attempts to obtain long-term repopulation from EBs in earlier studies. Failure to observe robust hematopoietic reconstitution directly from EBs led several groups to turn to alternative protocols employing coculture with supportive cell lines. PgP-1+ CD44+ Lineage− cells purified from ESC and cocultured with a bone marrow stromal cell line, cytokines, and fetal

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liver–conditioned medium were able to restore the hematopoietic compartment of primary and secondary recipients (Palacios et al., 1995). However, this report has not yet been replicated in the literature. Other work found that ESC cultured directly with OP9 stroma could generate lymphoid populations (Nakano et al., 1994). This observation, along with data showing that HoxB4 overexpression could expand bone marrow progenitors in a nontransforming manner, led to the discovery that retroviral transduction of EBs with HoxB4, followed by OP9 stroma coculture, is a robust and reproducible protocol for the specification and expansion of ESC-HSC from mESC (Kyba et al., 2002; Wang et al., 2005). ESCHSC derived in this system are capable of long-term, serial, multilineage repopulation of mice, demonstrated convincingly via retroviral marking studies (Wang et al., 2005), although the requirement for ectopic gene expression clearly shows that key signals required for the full developmental maturation of ESC-HSC are absent in this system. However, progress has recently been made towards the identification of morphogens that might be exploited to usurp the requirement for ectopic gene expression. BMP-4 was recently shown to induce the specification of blood progenitors in differentiating ESC via the Cdx-Hox pathway (Lengerke et al., 2007). Also, the large volume of cells required to rescue recipient animals from lethal irradiation, as well as the heterogeneous nature of the repopulating cell population, highlights the need to identify and purify the ESC-HSC. This work is currently underway.

Critical Parameters and Troubleshooting The following are the key variables in guaranteeing the successful application of this protocol. Maintain healthy OP9 stromal cell cultures Optimally maintained OP9 stromal cells should grow slowly (passage every ∼4 to 5 days at a split ratio of 1:3), should never be grown to more than 80% confluence, and should not be maintained in culture >4 weeks. If OP9 cultures begin to grow too slowly, it may be necessary to screen and select serum lots. Always use FRESHLY prepared 10% IMDM + cytokine cocktail Do not prepare a large volume of 10% IMDM + cytokine cocktail prior to the day

of actual use. Always prepare the medium on the day of infecting or passaging the culture. If using recipient animals that weigh >22 g or a different experimental model (i.e., mouse stains other than Rag-2−/− γc−/− ), the optimal irradiation dose MUST be determined experimentally. Careful experimentation has demonstrated that ESC-HSC are not identical to bone marrow HSC in their ability to mediate the rescue of mice from lethal irradiation. Whereas bone marrow HSC can readily rescue a Rag2−/− γc−/− mouse from 11 gy of irradiation, ESC-HSC cannot. Similarly, ESC-HSC do not compete well with radio-resistant host cells when transplanted into mice subjected to <8.5 gy of irradiation. Thus, 9.25 gy was found to be the optimal irradiation dose for the transplantation of Rag-2−/− γc−/− recipient animals with ESC-HSC in mice weighing weighing 15 to 22 g. If a different strain or size of mouse is going to be transplanted, the optimal irradiation dose must be determined experimentally to guarantee success and prevent self rescue. Always verify titer of MSCV-HoxB4-IRES-GFP virus Do not simply prepare supernatant and then use a volume of unknown titer for these experiments. Always determine the titer experimentally (i.e., using 3T3 cells; see Pear, 1996) and then perform infection with an MOI of 5 to 10. Verify adequate levels of hematopoietic specification in differentiating EBs If expansion of infected cells on OP9 stroma is not observed, verify that hematopoietic specification is occurring in differentiating EBs or purify hematopoietic progenitors from differentiating EBs (CD41+ c-kit+ ) via flow cytometry and infect only this population. It has been shown experimentally that the CD41+ c-kit+ compartment of day 6 EBs contains all hematopoietic progenitors and is the population that expands on OP9 stroma in response to HoxB4 infection. Adequate hematopoietic specification in differentiating EBs can be verified by examining the frequency of CD41+ c-kit+ cells by flow cytometry (2% to 5% expected in day 6 EBs; highly variable between ESC lines), observing a strong upregulation of runx1 or scl expression via RT-PCR between day 4 and day 6 of EB differentiation, or plating 200,000 to 500,000 day 6 EB-derived cells in 3 ml methylcellulose supplemented with hematopoiesis-promoting cytokines (M3434,

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StemCell Technologies) and observing the appearance of differentiated mixed colonies. If you fail to see evidence of the specification of hematopoietic progenitors, or secondary embryoid bodies dominate the CFU assay, then it may be necessary to screen lots of differentiation serum or to subclone an ESC line with good baseline hematopoietic differentiation activity.

Anticipated Results This protocol should result in the generation of a large number of OP9-expanded cells (160–200 × 106 ) and result in the reconstitution of >90% of the peripheral blood of recipient animals.

Time Considerations From beginning to end, this protocol takes 16 days before the cells are ready for transplantation. EBs differentiate for 6 days prior to infection with virus, and infected cells expand on OP9 stroma for at least 10 days prior to transplantation into recipients mice but can be allowed to expand for up to 12 days without compromising high-level hematopoietic reconstitution.

Literature Cited Brent, L., Sherwood, R.A., Linch, D.C., and Gale, R.E. 1990. Failure of embryonic mouse cells to engraft in immunocompetent allogeneic recipients. Br. J. Haematol. 74:549-551. Burt, R.K., Verda, L., Kim, D.A., Oyama, Y., Luo, K., and Link, C. 2004. Embryonic stem cells as an alternate marrow donor source: Engraftment without graft-versus-host disease. J. Exp. Med. 199:895-904. Cepko, C. 1996. Preparation of a specific retrovirus producer cell line. Curr. Protoc. Mol. Biol. 63:9.10.1-9.10.13. Chen, U., Kosco, M., and Staerz, U. 1992. Establishment and characterization of lymphoid and myeloid mixed-cell populations from mouse late embryoid bodies, “embryonic-stem-cell fetuses”. Proc. Natl. Acad. Sci. U.S.A. 89:25412545.

Keller, G. 2005. Embryonic stem cell differentiation: Emergence of a new era in biology and medicine. Genes Dev. 19:1129-1155. Kyba, M., Perlingeiro, R.C., and Daley, G.Q. 2002. HoxB4 confers definitive lymphoid-myeloid engraftment potential on embryonic stem cell and yolk sac hematopoietic progenitors. Cell 109:29-37. Lengerke, C., Schmitt, S., Bowman, T.V., Jang, I.H., Maouche-Chretien, L., McKinney-Freeman, S., Davidson, A.J., Hammerschmidt, M., Rentzsch, F., Green, J.B.A., Zon, L.I., and Daley, G.Q. 2008. BMP and Wnt specify hematopoietic fate by activation of the Cdx-Hox pathway. Cell Stem Cell 2:72-82. Muller, A.M. and Dzierzak, E.A. 1993. ES cells have only a limited lymphopoietic potential after adoptive transfer into mouse recipients. Development 118:1343-1351. Nakano, T., Kodama, H., and Honjo, T. 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265:10981101. Palacios, R., Golunski, E., and Samaridis, J. 1995. In vitro generation of hematopoietic stem cells from an embryonic stem cell line. Proc. Natl. Acad. Sci. U.S.A. 92:7530-7534. Pear, W. 1996. Transient transfection methods for preparation of high-titer retroviral supernatants. Curr. Protoc. Mol. Biol. 68:9.11.1-9.11.18. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. (eds.) 2008. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Sauvageau, G., Thorsteinsdottir, U., Eaves, C.J., Lawrence, H.J., Largman, C., Lansdorp, P.M., and Humphries, R.K. 1995. Overexpression of HoxB4 in hematopoietic cells causes the selective expansion of more primitive populations in vitro and in vivo. Genes Dev. 9:1753-1765. Verda, L., An Kim, D., Ikehara, S., Statkute, L., Bronesky, D., Petrenko, Y., Oyama, Y., He, X., Link, C., Vahanian, N.N., and Burt, R.K. 2007. Hematopoietic mixed chimerism derived from allogeneic embryonic stem cells prevents autoimmune diabetes mellitus in NOD mice. Stem Cells [Epub. Nov. 1, 2007]. Wang, Y., Yates, F., Naveiras, O., Ernst, P., and Daley, G.Q. 2005. Embryonic stem cell-derived hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 102:19081-19086.

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Differentiation of Mouse Embryonic Stem Cells into Blood

UNIT 1F.4

Yunglin D. Ma,1, 2 Jesse J. Lugus,1, 3 Changwon Park,1 and Kyunghee Choi1, 2, 3 1

Department of Pathology and Immunology, Washington University School of Medicine, St. Louis, Missouri 2 Developmental Biology Program, Washington University School of Medicine, St. Louis, Missouri 3 Molecular Cell Biology Program, Washington University School of Medicine, St. Louis, Missouri

ABSTRACT Embryonic stem (ES) cells can be maintained as pluripotent stem cells or induced to differentiate into many different somatic cell types. As ES-derived somatic cells can potentially be used for cell transplantation or cell-based therapy, ES cells have gained much scientific and general public attention. Successful derivation of blood from ES cells for tissue engineering will require a comprehensive understanding of inductive signals and downstream effectors involved in blood lineage development. Ideally, directed differentiation of ES cells into blood and isolation of pure hematopoietic progenitors will enhance our ability to utilize ES-derived blood cells for future clinical applications. The protocols provided in this unit describe methods of maintaining and differentiating mouse ES cells as well as identifying and isolating hematopoietic progenitors by utilizing flow C 2008 cytometry and progenitor assays. Curr. Protoc. Stem Cell Biol. 6:1F.4.1-1F.4.19.  by John Wiley & Sons, Inc. Keywords: embryonic stem cells r ES cells r in vitro differentiation r hematopoietic progenitors

INTRODUCTION The study of early events in hematopoietic development and the genes active during this process have been of great interest to investigators since the early 1900s. The modern tools of molecular biology and genetics, notably the development of genetic null animals through targeted gene deletion, have been utilized with great success to determine the individual genes and gene products required for discrete stages of hematopoietic development. Complicating this notion is the underlying importance of an intact hematopoietic system for mammalian development. Specifically, abrogation of a number of hematopoietic genes, including Gata1, Scl, Gata2, and Runx1, lead to embryonic lethality and preclude further analysis of animal tissues (Pevny et al., 1991; Tsai et al., 1994; Shivdasani et al., 1995; Okuda et al., 1996; Wang et al., 1996). Importantly, the use of embryonic stem (ES) cell–derived blood cells has allowed researchers to circumvent early lethality as well as generate large numbers of specific types of cells for further genetic and biochemical analyses. To this end, it has become highly desirable for researchers to master the technique of generating blood cells of different lineages in vitro from ES cells carrying targeted mutations. Additionally, recent work using transgenic ES cells, combined with a further understanding of the signaling and transcriptional requirements for hematopoiesis, has led to the ability to augment gene dosage during specific temporal periods and to understand the role of specific soluble factors in the development of cell types of interest. Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1F.4.1-1F.4.19 Published online July 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f04s6 C 2008 John Wiley & Sons, Inc. Copyright 

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This unit contains protocols for in vitro differentiation of mouse ES cells to blood lineages in serum-containing (Basic Protocol 1) and serum-free (Alternate Protocol 1) conditions, assessing the differentiated cells by flow cytometry (Basic Protocol 2), sorting cells (Alternate Protocol 2), preparing MEF feeder cells (Support Protocol 1), and maintaining ES cells (Support Protocol 2). NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. BASIC PROTOCOL 1

IN VITRO DIFFERENTIATION OF MOUSE ES CELLS TO BLOOD LINEAGES IN THE PRESENCE OF SERUM Many different methods of in vitro differentiation of ES cells can efficiently be used to generate the progeny of all three primary germ layers—endoderm, ectoderm, and mesoderm. The most typical method is to differentiate ES cells in a stromal cell–independent manner to give rise to three-dimensional, differentiated cell masses called embryoid bodies (EBs; Wiles and Keller, 1991; Park et al., 2004). The following protocol describes a method for differentiating ES cells to three-dimensional EBs in serum; differentiation in serum-free medium is described in Alternate Protocol 1. In the following protocol, the fetal bovine serum (FBS) contains all of the necessary growth factors. Additional growth factors (e.g., kit ligand and IL-3) are added to media (see Reagents and Solutions) to ensure healthy EB formation.

Materials Mouse ES cells, 3rd passage or more (up to 5 to 6 passages) after thawing (see Support Protocol 2) ES-IMDM medium (see recipe) 0.25% trypsin/EDTA (see recipe) Fetal bovine serum (FBS) for differentiation (see recipe) IMDM medium (see recipe) IMDM medium (see recipe) containing 10% (v/v) FBS for differentiation 2% (w/v) eosin in phosphate-buffered saline (PBS; see recipe) Serum differentiation medium, liquid (see recipe for ES differentiation media) 25-cm2 tissue culture flasks (Techno Plastic Product AG, cat. no. 90026; http://www.tpp.ch/), gelatinized (see Support Protocol 2, step 1) 14-ml polypropylene round-bottom snap-cap tubes (Becton Dickinson, cat. no. 352059) Hemacytometer 100-mm bacterial petri dishes (Kord-Valmark cat. no. 900; http://www.kord-valmark.com/) Centrifuge (e.g., Sorvall model RT7-RTH250) Additional reagents and equipment for maintaining ES cells (Support Protocol 2) Set up cultures 1. At a time point 2 days prior to setting up differentiation, split ES cells, seeding 4 × 105 ES cells per gelatinized 25-cm2 flask into 6 ml ES-IMDM medium. 2. Change medium the next day. Differentiation of mESCs to Blood

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Set up differentiation 3. Aspirate the medium from the flask. 4. Add 1 ml of 0.25% trypsin/EDTA at room temperature, swirl, and remove quickly. This wash can also be performed using PBS, but a preliminary wash with trypsin/EDTA prior to trypsinization in step 5 seems to be more effective in dispersing ES clumps into single cells.

5. Add 1 ml of 0.25% trypsin/EDTA at room temperature and wait until cells start to detach. This usually takes ∼10 to 30 sec. Do not over-trypsinize cells.

6. Stop the reaction by adding 1 ml of FBS (the same lot that is to be used for differentiation) and 4 ml of IMDM medium at room temperature and pipetting up and down to make a single-cell suspension. Transfer to a 14-ml snap-cap tube. It is important not to have cell clumps.

7. Centrifuge 5 to 10 min at 170 × g, room temperature. Aspirate the supernatant. 8. Wash the cell pellet by adding 10 ml of IMDM without FBS. Centrifuge 5 to 10 min at 170 × g, room temperature. Aspirate the supernatant. 9. Resuspend the cell pellet in 6 ml of IMDM with 10% FBS (for differentiation) and count viable ES cells in an aliquot using a 2% eosin solution in PBS and a hemacytometer. Make sure to count live ES cells only. For cell counting, eosin will stain the dead cells red; do not count red cells. After counting the cells, it is not necessary to recentrifuge the remaining cell suspension.

10. Plate cells in suspension culture in non-gelatinized 100-mm bacterial petri dishes (accommodating 10 ml medium) for differentiation into EBs. The cell number added for different EB stages is obtained empirically. We find that the culture becomes confluent and the EB differentiation becomes inefficient when more cells are seeded than recommended.

a. Add 6000 to 10,000 ES cells per ml of serum differentiation medium to obtain day-2.75 to day-3 EBs. Typically, blast colony forming cells (BL-CFCs) are measured between days 2.75 and 3. b. Add 4000 to 5000 cells per ml serum differentiation medium to obtain day-4 to day-5 EBs. Primitive erythroid progenitors are measured between days 4 and 5. c. Add ∼2000 cells per ml serum differentiation medium to obtain day-6 EBs. Definitive hematopoietic progenitors are measured between days 6 and 10. d. Add ∼500 cells per ml of methylcellulose medium (also see annotation below) to obtain day-8 to day-10 EBs. The methylcellulose medium contains the same reagents as liquid serum differentiation medium, except that methylcellulose is added to 1% of the final volume. ES cells aggregate to form EBs, which grow bigger with time. For more discussion on growth of EBs in suspension culture, see UNIT 1D.2. Primary differentiation is set up based on the cell number required for subsequent experiments. Total EB cell numbers typically obtained are as follows: day 2.75, ∼0.5-1 × 106 EB cells/10 ml of differentiation medium; day 4, ∼2-3 × 106 EB cells/10 ml of differentiation medium; day 6, ∼3-5 × 106 EB cells/10 ml of differentiation medium. We recommend differentiating ES cells in methylcellulose medium (see Reagents and Solutions) for obtaining late EBs (>day 8). Methylcellulose medium is semisolid; EBs

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Supplement 6

formed under methylcellulose conditions are uniform in size, while EBs formed under liquid differentiation conditions can be different in size. To obtain the late (>8-day) EBs, use methylcellulose medium starting at the beginning of culture. The methylcellulose medium contains the same reagents as liquid differentiation medium, except that methylcellulose is added to 1% of the final volume. To prepare >day-6 EBs, feed cells on day 6 by adding an additional 4 to 6 ml methylcellulose medium (see recipe) containing 0.5% methylcellulose instead of 1%, plus 1% (v/v) kit ligand conditioned medium and 1% (v/v) IL-3 conditioned medium (see Reagents and Solutions).

11. Differentiate the cells to the desired day and harvest for analysis or experimentation. ALTERNATE PROTOCOL 1

IN VITRO DIFFERENTIATION OF MOUSE ES CELLS TO BLOOD LINEAGES IN SERUM-FREE MEDIUM Mouse ES cells can also be differentiated in vitro to blood lineages in the absence of serum.

Additional Materials (also see Basic Protocol 1) Serum replacement (SR, Invitrogen, cat. no. 10828-028) Serum-free differentiation medium (see recipe for ES differentiation media) PFHM-II (Protein-Free Hybridoma Medium; Invitrogen, cat. no. 12040-077) Set up cultures 1. At a time point 2 days prior to setting up differentiation, split ES cells, seeding 4 × 105 ES cells per gelatinized 25-cm2 flask into 6 ml ES-IMDM medium. 2. Change medium the next day.

Set up differentiation 3. Aspirate the medium from the flask. 4. Add 1 ml of 0.25% trypsin/EDTA at room temperature, swirl, and remove quickly. This wash can also be performed using PBS, but a preliminary wash with trypsin/EDTA prior to trypsinization in step 5 seems to be more effective in dispersing ES clumps into single cells.

5. Add 1 ml of 0.25% trypsin/EDTA at room temperature and wait until cells start to detach. This usually takes ∼10 to 30 sec. Do not over-trypsinize cells.

6. Stop the reaction by adding 1 ml of FBS (the same lot that is to be used for differentiation) and 4 ml of IMDM medium at room temperature and pipetting up and down to make a single-cell suspension. Transfer to a 14-ml snap-cap tube. It is important not to have cell clumps.

7. Centrifuge 5 to 10 min at 170 × g, room temperature. Aspirate supernatant. 8. Wash the cell pellet by adding 10 ml of IMDM without FBS. Centrifuge 5 to 10 min at 170 × g, room temperature. 9. Resuspend the cell pellet in 6 ml of IMDM with 10% serum replacement or just IMDM (serum free) and count viable ES cells using 2% eosin solution in PBS.

Differentiation of mESCs to Blood

Make sure to count live ES cells only. For cell counting, eosin will stain the dead cells red; do not count red cells. After counting the cells, it is not necessary to recentrifuge the remaining cell suspension.

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Current Protocols in Stem Cell Biology

10. Plate cells in suspension culture in non-gelatinized 100-mm bacterial petri dishes (accommodating 10 ml medium) for differentiation into EBs. a. Add 8000 to 10,000 ES cells per ml of serum-free differentiation medium to obtain day-2.75 to day-3 EBs. b. Add 6000 to 7000 cells per ml of serum-free differentiation medium to obtain day-4 to day-5 EBs. c. Add ∼6000 cells per ml of serum-free differentiation medium to obtain day-6 EBs. We find that ES differentiation in serum free conditions is less efficient, thus we add more cells. Normally, we set up serum-free differentiation conditions to examine early time points, i.e., up to day 6.

11. Differentiate the cells to the desired day and harvest for analysis or experimentation.

PREPARATION OF IRRADIATED MOUSE EMBRYONIC FIBROBLASTS (MEFs) FOR USE AS FEEDER CELLS

SUPPORT PROTOCOL 1

Materials Mitotically arrested MEFs (PMEF; Specialty Media) 0.25% trypsin/EDTA (see recipe) FBS for ES cell culture (see recipe) IMDM medium (see recipe) Freezing medium: 90% FBS (for ES culture)/10% DMSO Liquid nitrogen 175-cm2 tissue culture flasks, gelatinized (see Support Protocol 2, step 1) 50-ml centrifuge tubes Centrifuge Hemacytometer Cryovials Liquid nitrogen freezer Additional reagents and equipment for γ irradiation of MEFs (UNIT 1C.3) 1. Thaw one vial of MEF cells into four gelatinized 175-cm2 flasks and grow at 37◦ C in a 5% CO2 incubator. 2. When confluent, passage the cells once into sixteen gelatinized 175-cm2 flasks. 3. Once confluent, trypsinize and harvest cells as follows: a. Aspirate the medium from the flask. Add 3 ml of 0.25% trypsin/EDTA at room temperature, swirl, and remove quickly. b. Add 3 ml of 0.25% trypsin/EDTA at room temperature and wait until cells start to detach. c. Stop the reaction by adding 3 ml of FBS (for culture) and 4 ml of IMDM medium at room temperature. d. Pipet up and down to make a single-cell suspension. e. Collect cells from flasks into 50-ml centrifuge tubes. Since there are sixteen flasks, one will have three to four 50-ml tubes containing the cell suspension.

f. Centrifuge the 50-ml tubes for 5 to 10 min at 170 × g, room temperature. Aspirate supernatants, resuspend each cell pellet in 10 ml IMDM medium containing 10% FBS (for culture), and combine cells in one 50-ml tube. Current Protocols in Stem Cell Biology

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1F.4.5 Supplement 6

4. Irradiate cells with 5000 rad from a γ source (UNIT 1C.3). 5. After irradiation, centrifuge cells at 5 min at 170 × g, room temperature. Aspirate the medium. 6. Count cells using hemacytometer and resuspend the cell pellet in freezing medium (90% FBS/10% DMSO) at a density of 1-1.25 × 106 cells/ml. Divide into 1-ml aliquots in cryovials. These will be single-use aliquots appropriate for seeding a 25-cm2 flask in Support Protocol 2 (see Support Protocol 2, step 2). Normally, PMEFs are seeded at 50,000 cells/cm2 .

7. Store cells at −80◦ C overnight, then transfer to liquid nitrogen (−150◦ C or lower). SUPPORT PROTOCOL 2

MOUSE ES CELL MAINTENANCE Mouse ES cells grow rapidly, with an average division time of ∼8 hr. In the authors’ laboratory, we normally split ES cells every 2 days and do not keep ES cells in culture for a long time after the cells are thawed. Typically, a new vial of cells is thawed after the previous batch of cells have undergone 5 to 6 passages. We have found that ES cells maintained on feeder cells (irradiated mouse embryonic fibroblasts, MEFs; see Support Protocol 1) give consistent in vitro differentiation behavior. The following protocol describes how to maintain ES cells.

Materials 0.1% gelatin (see recipe) γ-irradiated MEFs (see Support Protocol 1) MEF medium (see recipe) ES cells, frozen, passage 12 to 18 ES-DMEM medium (see recipe) ES-IMDM medium (see recipe) 0.25% trypsin/EDTA (see recipe) ES cell freezing medium (see recipe) 25-cm2 tissue culture flasks (Techno Plastic Products AG cat. no. 90026; http://www.tpp.ch/) 14-ml polypropylene round-bottom tube (Becton Dickinson; cat. no. 352059) Centrifuge (e.g., Sorvall model RT7-RTH250) Hemacytometer Day 1 1. Gelatinize a 25-cm2 flask by adding 3 ml of 0.1% gelatin, swirling to cover the entire surface, and letting the flask sit at room temperature for 10 to 20 min. 2. Thaw a vial of γ-irradiated MEF cells (prepared as in Support Protocol 1) in a 37◦ C water bath and transfer cells to a 14-ml snap-cap tube. Specialty Media sells frozen aliquots of MEFs.

3. Add 9 ml of fresh MEF medium and centrifuge the cells for 5 min at 170 × g, room temperature. Aspirate the supernatant. Be extremely careful not to disturb the cell pellet.

4. Resuspend the cells in 6 ml of MEF medium.

Differentiation of mESCs to Blood

5. Aspirate gelatin solution from the flask prepared in step 1 and transfer the suspension of MEF cells from step 4 to the flask. Place the flask in a 37◦ C incubator with 5% CO2 . All subsequent culture will be in the 37◦ C incubator with 5% CO2 .

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We have also had great success with STO cells mitotically arrested with mitomycin C instead of γ -irradiation. Treat STO cells seeded the previous day at 50,000 cells/cm2 with mitomycin C (Sigma cat. no., M0503, 10 μg/ml in MEF medium) for 2 to 3 hr, wash, and feed with 6 ml fresh MEF medium. Add ES cells the next day.

Day 2 6. Thaw a vial of mouse ES cells in a 37◦ C water-bath and transfer cells to a 14-ml snap cap tube. We typically use passages between 12 and 18 for starting the ES culture.

7. Add 9 ml of fresh ES-DMEM medium and centrifuge the cells for 5 min at 170 × g, room temperature. Aspirate the supernatant. Be extremely careful not to disturb the cell pellet.

8. Resuspend ES cells in 6 ml of fresh ES-DMEM medium. Remove MEF medium from the 25-cm2 flask containing feeder cells and transfer ES cells to the flask.

Day 3 9. Feed cells with 6 ml ES-DMEM medium. Prepare a new gelatin-coated 25-cm2 flask and thaw out MEFs as in step 1. Day 4 or 5 10. Split ES cells and passage onto MEFs. Aspirate medium and wash briefly with 1 ml of 0.25% trypsin/EDTA. After trypsin/EDTA has been removed, add another 1 ml of fresh 0.25% trypsin/EDTA and place the flask in a 37◦ C incubator for 10 to 30 sec, enough time for the cells to lift off the flask. Depending on the number of viable ES cells recovered from a vial of freshly thawed cells, you may need additional 1 or 2 days in culture before the growth of the newly thawed cells is sufficient. Ideally, a given flask will contain a large number of smaller colonies rather than a very small number of large colonies. Passaging at the appropriate time will prevent the latter scenario.

11. Add 5 ml of ES-DMEM and pipet up and down to break up the cell clumps. Transfer to a 14-ml snap-cap tube and centrifuge for 5 min at 170 × g, room temperature. 12. Aspirate the supernatant and resuspend the cell pellet in 6 ml of fresh ES-DMEM medium. 13. Count cells using a hemacytometer, being careful to distinguish between ES cells and feeder cells. ES cells are smaller, translucent and uniform in cell size, while mitotically inhibited MEFs are much bigger and granular.

14. Plate 8 × 105 ES cells in a new 25-cm2 flask with MEFs in 6 ml ES-DMEM.

Second and subsequent passages 15. Day after the 1st passage: Feed ES cells with 6 ml fresh ES-DMEM and prepare a new 25-cm2 flask of MEFs as described above. 16. 2 days after the 1st passage: Passage cells again as in steps 10 to 14. The authors of this unit do not prepare cells to generate EBs after the first passage, as the ES cells do not differentiate well. We passage cells a minimum of twice after thawing before preparing cells to differentiate into EBs.

17. Day after the 2nd passage: Feed cells with 6 ml fresh ES-DMEM and prepare new 25-cm2 flask with MEFs. During the 1st and 2nd passages after thawing, the ES cells can be frozen at a density of 2 × 106 cells/ml in ES cell freezing medium.

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18. 2 days after the 2nd passage: Passage again, both onto MEFs to maintain the ES line and also onto a gelatinized flask with ES-IMDM to prepare ES cells for in vitro differentiation: a. Gelatinize a 25-cm2 flask by adding 3 ml of 0.1% gelatin, swirling to cover the entire surface, and letting the flask sit at room temperature for 10 to 20 min. b. After incubation, remove gelatin and add 6 ml of ES-IMDM. Plate 4 × 105 ES cells in this flask. c. Also, plate 8 × 105 ES cells onto a 25-cm2 flask of MEFs, as described above. This makes a 3rd passage. The authors normally prepare one 25-cm2 flask of ES cells for in vitro differentiation. One can prepare more than one 25-cm2 flask of cells depending on the scale of in vitro differentiation.

19. Day after the 3rd passage: Feed cells on MEFs and on a gelatinized flask with 6 ml of ES-DMEM or ES-IMDM, respectively. Prepare a new flask of MEFs as described above. 20. 2 days after the 3rd passage: Passage cells on MEFs as and onto a gelatin-coated flask as described in step 18. Use cells on a gelatinized flask from the 3rd passage to differentiate into EBs (Basic Protocol 1 or Alternate Protocol 1). 21. Repeat steps 15 and 16 until cells have been passaged 5 to 6 times, then discard cells. We find that ES cells do not differentiate well in culture after 5 to 6 passages. BASIC PROTOCOL 2

FLOW CYTOMETRIC ANALYSIS OF EB CELLS Hematopoietic progenitors present within EBs can be assayed by flow cytometry. A flow cytometer or fluorescence activated cell sorter (FACS) utilizes cells treated with monoclonal antibodies, conjugated to different fluorochromes, against cell surface proteins or intracellular markers to identify, analyze, and isolate specific EB cell population (Chung et al., 2002; Lugus et al., 2007). The authors of this unit typically use day 2 to 3 EBs to analyze mesoderm (FLK1+ ) by utilizing a FLK1 monoclonal antibody, and day-4 to day-8 EBs to analyze hematopoietic (CD45+ and TER119+ ) and endothelial (CD31+ and VEcadherin+ ) progenitors. See Table 1F.4.1 for antibodies used to characterize progenitors.

Materials Mouse EBs (Basic Protocol 1 and Alternate Protocol 1) 7.5 mM EDTA (BioRad, cat. no. 161-0729) in PBS, pH 7.4 (see recipe) IMDM medium (see recipe) Washing buffer: 4% (v/v) FBS (for culture) in PBS (see recipe) Primary antibody at appropriate dilution (Table 1F.4.1) in washing buffer Secondary antibody (if needed) at appropriate dilution (Table 1F.4.1) in washing buffer

Differentiation of mESCs to Blood

50-ml centrifuge tubes (Fisher, cat. no. 14-432-22) Centrifuge with microtiter plate carrier 20-G needle (Fisher, cat. no. 14826-5C) Hemacytometer 96-well plate with V-bottom wells (Fisher, cat. no. 07-200-96) 5-ml polypropylene tubes (VWR, cat. no. 60818-500) CellQuest (Becton-Dickinson) or FlowJo (Tree Star, Inc., http://www.treestar.com) software FACScan or FACSCalibur flow cytometer (BD Biosciences) Additional reagents and equipment for flow cytometry (Robinson et al., 2008)

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Table 1F.4.1 Antibodies Used to Characterize Mesoderm, Endoderm, and Hematopoietic Progenitors

Target antigena

Primary antibodyb

Secondary antibodyb

FLK1 (VEGF-R2, Ly-73)

PE-conjugated rat anti-mouse FLK1 (1:200; BD Pharmingen, cat. no. 555308)

—

CD45 (Ly-5)

FITC-conjugated anti-mouse CD45 (1:200; eBioscience, cat. no. 11-0451)

—

TER119 (Ly-76)

FITC-conjugated anti-mouse TER119 (1:200; eBioscience, cat. no. 11-5921)

—

CD31 (PECAM-1)

PE-conjugated rat anti-mouse CD31 (1:200; BD Pharmingen, cat. no. 553373)

—

VE-Cadherin (CD144, Cadherin-5)

Purified rat anti-mouse Alexa Fluor 488 goat anti-rat CD144 (1:500; BD IgG(H+L) (1:1000; Invitrogen, Pharmingen, cat. no. 555289) cat. no. A11006)

a Alternative names appear in parentheses. b Dilution and supplier appear in parentheses.

Dissociate EBs into single-cell suspension 1. Collect EBs in a 50-ml tube and centrifuge 1 min at 170 × g, room temperature, or by letting them settle at room temperature for 10 to 20 min. 2. Remove the supernatant and treat EBs with 1 ml of 7.5 mM EDTA /PBS (pH 7.4) for 1 min at 37◦ C. Trypsin/EDTA can be used to dissociate EB cells when FLK1 is the only antigen to be analyzed, as we find that FLK1 is resistant to trypsin/EDTA treatment.

3. Add 9 ml of IMDM to dilute EDTA. Vortex quickly and centrifuge the cells for 5 min at 170 × g, room temperature. EDTA should be removed as soon as possible to minimize the exposure time to EDTA. Prolonged exposure to EDTA can lead to cell death.

4. Aspirate the supernatant and add 3 ml of washing buffer. 5. Pass through a 20-G needle 4 to 5 times to generate a single-cell suspension, and count viable cells with 2% eosin in PBS. Make sure to count live ES cells only. For cell counting, eosin will stain the dead cells red; do not count red cells.

6. Centrifuge the cells for 5 min at 170 × g, room temperature. After centrifugation, aspirate the supernatant and resuspend the cells at a density of 5 × 106 cells/ml in washing buffer.

Stain cells 7. Place cells into individual wells of a V-shaped 96-well plate at 5 × 105 cells/well. Centrifuge the plate 5 min at 170 × g, room temperature. 8. Aspirate the supernatants from the wells of the 96-well plate using a multichannel pipettor. Add primary antibody at an appropriate dilution in 100 μl of washing buffer. Incubate on ice (or 4◦ C) for 15 min. If your primary antibody is directly conjugated to a fluorochrome, you can skip the secondary antibody staining and continue with step 13. Current Protocols in Stem Cell Biology

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1F.4.9 Supplement 6

9. After incubation, add additional 100 μl of washing buffer to each well and centrifuge cells at 170 × g, room temperature, for 5 min. Remove the supernatant. 10. Wash cells in 150 μl per well of washing buffer and remove the supernatant. 11. Repeat step 10 for a total of three washes. 12. After three washes, add 100 μl of freshly diluted secondary antibody in staining buffer and incubate on ice (or 4◦ C) for 15 min in the dark. The plate has to be kept in the dark if the secondary antibody is directly conjugated to fluorochrome.

Perform flow cytometry and analyze data 13. After incubation, wash cells three times as in steps 9 to 11, resuspend in 150 μl of washing buffer, and transfer to a 5-ml polypropylene tube for flow cytometric analysis. 14. Acquire flow cytometric data on a FACScan or FACSCalibur flow cytometer (Robinson et al., 2008) and analyze with CellQuest or FlowJo software. ALTERNATE PROTOCOL 2

CELL SORTING AND IN VITRO CULTURE OF SORTED CELL POPULATIONS The staining for cell sorting is performed the same way as for flow cytometric analysis (Basic Protocol 2). The variant steps are described below.

Additional Materials (also see Basic Protocol 2) 40-μm nylon-mesh cell strainer (BD Falcon, cat. no. 352340) MoFlo cell sorter (BD Biosciences) 14-ml tubes (Fisher, no. 14-959-49B) 1. Prior to sorting, filter stained cells through a 40-μm nylon-mesh cell strainer. 2. Sort cells using the MoFlo cell sorter into a 14-ml tube (Fisher, no. 14-959-49B) containing 2 ml of FBS (for culture). Reanalyze the sorted cells for the same antigens as used for sorting on a FACSCalibur flow cytometer to determine the sorting efficiency. EB cells are notorious for their stickiness. For pure cell sorting with good yields, the sample must be as close to an absolute single-cell suspension as possible.

3. Use sorted cells for hematopoietic progenitor assays (Basic Protocol 3). Alternatively, use the cells to make RNA for gene-expression studies. BASIC PROTOCOL 3

HEMATOPOIETIC PROGENITOR ASSAYS Hematopoietic progenitors present in EBs can also be assayed by directly replating EB cells. Day-2.75 to day-3 EBs are typically used for blast colony assay (Kennedy et al., 1997; Choi et al., 1998), day-4 EBs for primitive erythroid colony assay, and day-6 to day-10 EBs for definitive erythroid and myeloid progenitor analyses (Wiles and Keller, 1991; Keller et al., 1993).

Materials

Differentiation of mESCs to Blood

Mouse EBs (Basic Protocol 1 or Alternate Protocol 1) IMDM medium (see recipe) 2× cellulase solution (see recipe) 0.25% trypsin/EDTA (see recipe) Collagenase solution (optional; for older EBs; see recipe)

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FBS (for differentiation; see recipe) IMDM medium (see recipe) containing 10% (v/v) FBS (for differentiation) 2% (w/v) eosin in phosphate-buffered saline (PBS; see recipe) Methylcellulose mixes for progenitor cells of interest (see recipe) 50-ml polypropylene conical tube (Becton Dickinson; cat. no. 352070) 20-G and 16- or 18-G needles with 3-ml syringes 14-ml polypropylene round-bottom tube (Becton Dickinson; cat. no. 352059) 35-mm and 150-mm bacterial dishes (Becton Dickinson; cat. no. 351008 and 351058, respectively) Inverted microscope Harvest EBs 1a. For EBs in liquid: Transfer medium containing EBs into a 50-ml tube. Wash the plate with IMDM and add to the 50-ml tube and let sit at room temperature for ∼10 to 20 min. EBs will settle to the bottom of the tube.

1b. For EBs in methylcellulose: Add an equal volume of 2× (2 U/ml) cellulase (final 1 U/ml) to EBs growing in methylcellulose and incubate 20 min at 37◦ C. Collect EBs in a 50-ml tube. Wash the plate with 10 ml IMDM. Add the wash to the tube of cells and allow the cells to settle 10 to 20 min at room temperature. 2. Aspirate the medium, add 3 ml of 0.25% trypsin/EDTA, and incubate for 3 min in a 37◦ C water bath. Use collagenase for older EBs (>day 8, for example). When collagenase is used, incubate EBs for 1 hr at 37◦ C.

Dissociate EBs 3. Vortex quickly and add 1 ml of FBS (for differentiation). Dissociate cells by passing through a 20-G needle 4 to 5 times. 4. Transfer to a 14-ml snap cap tube and centrifuge for 5 to 10 min at 170 × g, room temperature. Discard the supernatant. 5. Resuspend the cell pellet in 0.3 to 1 ml of IMDM containing 10% FBS (for differentiation). 6. Count the viable cells in an aliquot with 2% eosin in PBS. Make sure to count live ES cells only. For cell counting, eosin will stain the dead cells red; do not count red cells. At this point, there should be no cell clumps.

Culture cells 7. Add cell suspension to a 14-ml snap-cap tube containing methylcellulose mix corresponding to the cell of interest (see Reagents and Solutions). Vortex thoroughly and let it sit at room temperature for 5 to 10 min. Typically, the cells are used at 3-6 × 104 EB cells per 1 ml of methylcellulose medium.

8. Prepare 4 ml of methylcellulose mix for each of three replica dishes (35-mm bacterial petri dishes) for each sample. Using a syringe with a 16-G or 18-G needle, add 1 ml of the methylcellulose mix to each dish. Spread the methylcellulose mixture by gently tapping. The reason for making 4 ml of methylcellulose medium for each sample is that the methylcellulose medium is very viscous. The recipes for the blast colony, primitive erythroid colony, and definitive erythroid/myeloid progenitor assays are shown below.

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9. Put several 35-mm bacterial dishes (up to 6 dishes) into a 150-mm bacterial dish with a 35-mm open dish containing sterile water in the middle. Culture in a 37◦ C CO2 incubator. 10. Count colonies under inverted microscope 4 to 7 days after replating. Blast colonies develop from day-2.75 to day-3 EBs and contain cells with undifferentiated or blast morphology. Only blast colonies and secondary EBs form from day-2.75 to day-3 EBs. Secondary EBs are compact, and no individual cells can be identified; thus they can be easily distinguished from blast colonies. Primitive erythroid colonies developing from day-4 EBs are small and compact. Macrophage colonies developing from day-6 to day-8 EBs contain larger cells with granules. E-Mac colonies contain both erythroid cells and macrophages. Additional information on hematopoietic colonies is given in the original papers (Wiles and Keller, 1991; Keller et al., 1993; Kennedy et al., 1997; Choi et al., 1998).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Ascorbic acid Dissolve ascorbic acid (Sigma; cat. no. A-4544) at 5 mg/ml in autoclaved water and filter sterilize using a 0.22-μm filter. Prepare ascorbic acid solution fresh each time differentiation is set up.

Cellulase solution, 2× Dissolve cellulase (Sigma; cat. no. C-1794) in PBS (see recipe) at 2 U/ml. Filter sterilize through 0.45-μm filter. Store up to 1 to 2 months at −20◦ C.

Collagenase solution Dissolve 1 g of collagenase (Sigma; cat. no. C-0310) in 320 ml of PBS (see recipe). Filter sterilize through a 0.45-μm filter, then add 80 ml of FBS (for differentiation; see recipe). Divide into 50-ml aliquots and keep at –20◦ C up to 1 to 2 months.

D4T conditioned medium (CM) Seed D4T endothelial cells (Kennedy et al., 1997; Choi et al., 1998) in IMDM medium (see recipe) containing 10% FBS at a density of 25,000 cells/cm2 and begin incubation. When culture becomes 80% confluent, remove medium and replace with IMDM containing 4% FBS. Culture an additional 72 to 96 hr, and collect the supernatant. Centrifuge for 5 min at 170 × g, room temperature, to remove cell debris, then filter sterilize the supernatant utilizing a 0.45-μm filter unit. Divide into 10-ml aliquots and store at –80◦ C. Once thawed, D4T CM is kept at 4◦ C for about 1 week.

DMEM medium Dissolve 1 package of Dulbecco’s Modified Eagle Medium (DMEM) powder (Invitrogen, cat. no. 12100-046) in ∼800 ml autoclaved distilled water. Add 3.024 g NaHCO3 (Sigma, cat. no. S-5761), 10 ml penicillin/streptomycin (10,000 U; Invitrogen, cat. no. 6005140PG), and 25 ml of 1 M HEPES (Invitrogen, cat. no. 380-5630 PG). Bring up to 1 liter with autoclaved distilled water, filter through 0.22-μm filter, and store at 4◦ C up to 1to 2 months. Differentiation of mESCs to Blood

We normally use distilled water from Millipore Milli-Q purification system (QTUM000EX).

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Current Protocols in Stem Cell Biology

ES cell freezing medium 90% FBS (for culture; see recipe) with 10% DMSO (Sigma, cat. no. D-2650). ES cells are frozen at a density of 2–3 × 106 cells per ml of freezing medium. Add 1 ml of cells to each freezing vial. Store cells at −80◦ C overnight before transferring them to liquid nitrogen (less than −150◦ C).

ES differentiation media See Table 1F.4.2 for the composition of various ES differentiation media.

ES-DMEM medium Dulbecco’s modified Eagle medium (DMEM; Invitrogen, cat. no. 12100-046) containing: 15% (v/v) FBS (preselected for culture; see recipe) 2% (v/v) LIF (leukemia inhibitory factor) conditioned medium (see recipe) 1% (v/v) nonessential amino acids (Mediatech, cat. no. 25-025-CI) 1% (v/v) L-glutamine (Invitrogen, cat. no. 25030) 1.5 × 10−4 M MTG (see recipe) Store up to 1 month at 4◦ C ES-IMDM medium Iscove’s Modified Dulbecco’s Medium (IMDM; Invitrogen, cat. no. 12200-036) containing: 15% (v/v) FBS (preselected for culture; see recipe) 2% (v/v) LIF (leukemia inhibitory factor) conditioned medium (see recipe) 1% (v/v) nonessential amino acids (Mediatech, cat. no. 25-025-CI) 1% (v/v) L-glutamine (Invitrogen, cat. no. 25030) 1.5 × 10−4 M MTG (see recipe) Store up to 1 month at 4◦ C Table 1F.4.2 ES Differentiation Media Composition

Serum differentiation medium (liquid)

Serum differentiation medium (methylcellulose)

Serum-free differentiation medium (liquid)

2% methylcellulosea

—

55% (v/v)

—

FBS (preselected for differentiation)a

15% (v/v)

15% (v/v)

—

Serum replacementb

—

—

15% (v/v)

Ascorbic acid (5 mg/ml)a

50 μg/ml

50 μg/ml

50 μg/ml

L-glutamine (200 mM)c

2 mM

2 mM

2 mM

MTGa

To 4.5 × 10−4 M

To 4.5 × 10−4 M

To 4.5 × 10−4 M

—

—

5%

To 100%

To 100%

To 100%

PFHM-IId IMDM

e

a See recipe in Reagents and Solutions. b Invitrogen, cat. no. 10828-028. c Invitrogen, cat. no. 25030. d Protein-Free Hybridoma Medium; Invitrogen, cat. no. 12040-077. e Invitrogen, cat. no. 12200-036.

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Supplement 6

FBS for ES culture We normally prescreen FBS for ES culture. Typically, ES cells adapted to grow without feeder cells are used for testing serum. ES cells are maintained in test serum for 5 to 6 passages and scored for morphology; either differentiated or undifferentiated. A rapid, easy and quantifiable assessment of different FBS lots for use in ES cell propagation is to grow Oct4-GFP ES cells (Qi et al., 2004) in various lots of serum and perform flow cytometric analyses to assess GFP-positivity. A good lot of serum should maintain >95% of Oct4-GFP ES cells as GFP+ after 5 to 6 passages.

FBS for ES differentiation We normally prescreen FBS for ES differentiation. Typically, ES cells are differentiated in test serum and analyzed by flow cytometry for FLK1 staining or hematopoietic replating. A good lot of serum should generate ∼30% to 50% of FLK1+ cells when day-3 to day-4 EB cells are analyzed.

Gelatin, 0.1% Dissolve gelatin (Sigma G-1890) at 0.1% (w/v) in PBS (see recipe) and autoclave. Store up to 1 month at 4◦ C.

IL-3 IL-3 is from medium conditioned by X63 AG8-653 myeloma cells transfected with a vector expressing IL-3 (Genetics Institute, Inc., Cambridge, Massachusetts; Karasuyama and Melchers, 1988). IL-3-producing cells are seeded at 25,000 cells/cm2 and cultured in IMDM medium (see recipe) containing 10% FBS (for differentiation; see recipe). When culture becomes 80% confluent, remove medium and replace with IMDM containing 4% FBS (for differentiation). Culture an additional 72 to 96 hr, and collect the supernatant. Centrifuge for 5 min at 170 × g, room temperature, to remove cell debris, then filter sterilize the supernatant utilizing a 0.45-μm filter unit. Divide into 10-ml aliquots and store at –80◦ C. Once thawed, IL-3 CM may be kept at 4◦ C for ∼1 week.

IMDM medium Dissolve one package of Iscove’s Modified Dulbecco’s Medium (IMDM) powder (Invitrogen, cat. no. 12200-036) in ∼800 ml autoclaved distilled water. Add 3.024 g NaHCO3 (Sigma, cat. no. S-5761) and 10 ml penicillin/streptomycin (10,000 U; Invitrogen, cat. no. 6005140PG). Bring up to 1 liter with autoclaved distilled water, filter through 0.22-μm filter, and store at 4◦ C up to 1 to 2 months. We normally use distilled water from Millipore Milli-Q purification system (QTUM000EX).

Kit ligand

Differentiation of mESCs to Blood

Kit ligand (KC) is from medium conditioned by CHO cells transfected with a KL expression vector (Genetics Institute, Inc., Cambridge, Massachusetts). KLproducing cells are seeded at 25,000 cells/cm2 and cultured in IMDM medium (see recipe) containing 10% FBS (for differentiation). When culture becomes 80% confluent, remove medium and replace with IMDM containing 4% FBS (for differentiation). Culture an additional 72 to 96 hr, and collect the supernatant. Centrifuge for 5 min at 170 × g, room temperature, to remove cell debris, then filter sterilize the supernatant utilizing a 0.45-μm filter unit. Divide into 10-ml aliquots and store at –80◦ C. Once thawed, KL CM may be kept at 4◦ C for ∼1 week.

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LIF conditioned medium Chinese Hamster Ovary (CHO) cells transfected with the LIF (leukemia inhibitory factor) gene (Genetics Institute) are used as a source for LIF. Typically LIF is secreted from the cells into the medium at ∼5 μg/ml. LIF-producing cells are seeded at 25,000 cells/cm2 and cultured in IMDM medium (see recipe) containing 10% FBS (for culture). When culture becomes 80% confluent, remove medium and replace with DMEM containing 4% FBS (for culture). Culture an additional 72 to 96 hr, and collect the supernatant. Centrifuge for 5 min at 170 × g, room temperature, to remove cell debris, then filter sterilize the supernatant utilizing a 0.45-μm filter unit. Divide into 10-ml aliquots and store at –80◦ C. Once thawed, LIF CM may be kept at 4◦ C for about 1 week.

MEF medium Dulbecco’s modified Eagle medium (DMEM; Invitrogen, cat. no. 12100-046) containing: 15% (v/v) FBS (preselected for culture; see recipe) 1% (v/v) nonessential amino acids (Mediatech, Inc., cat. no. 25-025-CI) 1% (v/v) L-glutamine (Invitrogen, cat. no. 25030) 1.5 × 10−4 M MTG (see recipe) Store up to 1 month at 4◦ C Methylcellulose, 2% (w/v) Weigh a sterile 1-liter Erlenmeyer flask. Add ∼450 ml of sterile water. Bring to boil on a hot plate and keep boiling for 3 to 4 min. Add 20 g of methylcellulose (Fluka, cat. no. 64630), swirl quickly, and return the flask to the hot plate. Remove the flask quickly from the hot plate and swirl again when it starts to boil. Return the flask to the hot plate. Repeat three to four times. Weigh the flask with the solution, subtract the weight of the flask, and add sufficient sterile water (at room temperature) to make 500 ml of methylcellulose mixture. Let it sit on bench to cool down to room temperature. In a separate weighed flask, make 500 ml of 2× IMDM and filter sterilize (0.22-μm). Slowly add the 500 ml of 2× IMDM to the 500 ml of methylcellulose and mix vigorously. Put the mixture on ice until the medium becomes viscous. Make ∼100 ml aliquots and store frozen at –20◦ C. When ready to use, thaw and use a syringe to disperse methylcellulose (do not use pipets).

Methylcellulose mixes for progenitor assays See Table 1F.4.3 for the composition of the methylcellulose mixes for the progenitor assays.

Monothioglycerol (MTG), 1.5 × 10−4 M in medium Add the MTG by freshly diluting MTG (Sigma; cat. no. M-6145) 1:10 in DMEM and adding 12.4 μl per 100 ml of the medium to be prepared. Alternatively, β-mercaptoethanol (2-ME; Sigma, cat. no. M-7522) is used at 1 × 10−4 M. For a 100× stock solution adding 72 μl of 14 M 2-ME to 100 ml of PBS (see recipe). To use, add 1 ml per 100 ml of medium. Make sure that MTG or BME is made fresh.

Monothioglycerol (MTG), 4.5 × 10−4 M in medium Add the MTG by freshly diluting 26 μl of MTG into 2 ml of IMDM and adding 3 μl of this diluted MTG per ml of medium.

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Table 1F.4.3 Methylcellulose Mixes for Progenitor Assays

Blast

Primitive erythroid

Definitive erythroid and myeloid

2% methylcellulosea

55% (v/v)

55% (v/v)

55% (v/v)

FBS (preselected for differentiation)a

10% (v/v)

—

—

PDSb

—

10% (v/v)

10% (v/v)

12.5 μg/ml

12.5 μg/ml

12.5 μg/ml

2 mM

2 mM

2 mM

a

Ascorbic acid c

L-glutamine d

Transferrin

200 μg/ml

200 μg/ml −4

200 μg/ml −4

To 4.5 × 10

D4T conditioned mediuma

20% (v/v)

—

—

VEGFe

5 ng/ml

—

—

Kit ligand conditioned mediuma

1% (v/v)

—

1% (v/v)

EPOf

—

2 U/ml

2 U/ml

—

5% (v/v)

5% (v/v)

—

—

5 ng/ml

IL-3 conditioned mediuma

—

—

1% (v/v)

IL-6i

—

—

5 ng/ml

—

—

5-25 ng/ml

—

—

2-30 ng/ml

g

PFHM-II IL-1β

h

j

IL-11

k

G-CSF

l

M

—

—

3-5 ng/ml

m

—

—

2-5 ng/ml

n

To 100%

To 100%

To 100%

GM-CSF M-CSF IMDM

M

To 4.5 × 10

To 4.5 × 10−4 M

MTG

a

a See recipe in Reagents and Solutions. b Plasma-derived serum; Animal Technologies, http://www.animaltechnologies.com. c Invitrogen, cat. no. 25030. d Transferrin (Human) in IMDM (Boehringer-Mannheim/Roche, cat. no. 652202). e R&D Systems, cat. no. 293-VE. f Erythropoietin (Amgen Epogen NDC 55513-126-10). g Protein-Free Hybridoma Medium; Invitrogen, cat. no. 12040-077. h R&D Systems, cat no. 401-ML. i R&D Systems, cat. no. 406-ML. j R&D Systems, cat. no. 418-ML. k R&D Systems, cat. no. 414-CS. l R&D Systems, cat. no. 415-ML. m R&D Systems, cat. no. 416-ML. n Invitrogen, cat. no. 12200-036.

Phosphate-buffered saline (PBS), 10×

Differentiation of mESCs to Blood

Combine the following: 80 g NaCl 2 g KCl 14.4 g Na2 HPO4 2.4 g KH2 PO4 800 ml H2 O continued

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Heat to dissolve Adjust pH to 7.4 with HCl Add H2 O to 1 liter Filter sterilize through 0.22-μm filter Store up to 6 months at room temperature Trypsin/EDTA, 0.25% Dissolve 2.5 g of trypsin (Sigma, cat. no. T-4799) in 900 ml of PBS (see recipe). Add 2.16 ml of 0.5 M EDTA and bring up to 1 liter with PBS. Filter sterilize through 0.22-μm filter. Store in aliquots at –20◦ C. Once thawed, store at 4◦ C for up to 1 month.

COMMENTARY Background Information An alternate source of embryonic cells for studies of early embryonic events is the in vitro–differentiated progeny of ES cells. ES cells differentiate efficiently in vitro and give rise to three-dimensional, differentiated cell masses called embryoid bodies (EBs, reviewed in Park et al., 2005). ES cells can also be differentiated on stromal cells or type IV collagen without intermediate formation of the EB structure (Nakano et al., 1994; Nishikawa et al., 1998). Many different lineages have been reported to develop within EBs, including neuronal, muscle, endothelial, and hematopoietic lineages (reviewed in Park et al., 2005). Of these, the hematopoietic lineage has been the most extensively characterized. Hematopoietic progenitors develop sequentially within EBs. The first to develop is the Blast Colony-Forming Cell (BL-CFC). BL-CFCs are transient and develop prior to the primitive erythroid population (Choi et al., 1998; Lugus et al., 2007). Definitive erythroid and myeloid progenitors develop shortly after primitive erythroid progenitors. BL-CFCs form blast colonies in response to vascular endothelial growth factor (VEGF), a ligand for the receptor tyrosine kinase (FLK1). Gene expression analysis has indicated that cells within blast colonies (blast cells) express a number of genes common to both hematopoietic and endothelial lineages, including Scl, CD34, and Flk1 (Kennedy et al., 1997). In addition, blast cells are clonal and give rise to primitive and definitive hematopoietic as well as endothelial cells when replated in media containing both hematopoietic and endothelial cell growth factors (Kennedy et al., 1997; Choi et al., 1998). The developmental kinetics of various hematopoietic lineage precursors within EBs, as well as molecular and cellular studies of these cells, have demonstrated

that the sequence of events leading to the onset of hematopoiesis within EBs is similar to that found within the normal mouse embryo. In addition, EBs provide a large number of cells representing an early/primitive stage of development that is otherwise difficult to access in an embryo. Therefore, the in vitro differentiation model of ES cells is an ideal system for obtaining and studying primitive progenitors of all cell lineages.

Critical Parameters and Troubleshooting For ES cell maintenance and differentiation 1. The authors recommend that ES cells be healthy and fresh. Mouse ES cells grow rapidly, with an average division time of about 8 hr. Therefore, ES cells require frequent splitting. We normally split ES cells every 2 days and do not keep ES cells in culture for a long time after the cells are thawed. Typically, a new vial of cells is thawed after the initial cultures have undergone 5 to 6 passages. We recommend that ES cells be passed at least one time after the thaw, before setting up differentiation. We typically set up three independent differentiations from one thaw. 2. For ES cell differentiation, we add more cells for ES lines that differentiate poorly. 3. We find that liquid differentiation is good for obtaining early EBs (up to days 5 to 6) and methylcellulose differentiation for obtaining late EBs (days 6 to 14). The methylcellulose medium contains the same reagents as liquid differentiation medium, except that methylcellulose is added to 1% of the final volume. 4. Some maintain ES cells on gelatinized flasks without feeder cells. We find that ES cells maintained on feeder cells give more consistent in vitro differentiation results compared to those maintained on gelatinized flasks.

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5. When ES cells differentiate poorly, we check MTG and ascorbic acid. We typically open a new bottle of MTG every 1 to 2 months. Ascorbic acid needs to be made fresh every time a differentiation experiment is set up. 6. It is important to put only 4 × 105 ES cells per 25-cm2 flask, 2 days prior to differentiation. The ES cell confluency in ES-IMDM medium should not exceed 80%. ES cell differentiate poorly, if the cells are too confluent, but they also differentiate poorly if the culture is too sparse. 7. Pre-selected serum (see Reagents and Solutions) seems to be most critical for optimal generation of hematopoietic lineages.

to 3 differentiations, including hematopoietic replating and counting.

For replating 1. D4T conditioned medium (CM) appears to be important to obtain healthy blast colonies. D4T is an endothelial cell line which was generated from day-4 EB cells by infecting with retroviruses expressing polyoma middle T gene (Kennedy et al., 1997; Choi et al., 1998). We have not determined if other endothelial cell conditioned media will also support blast colony formation. 2. We typically use plasma-derived serum (PDS) for primitive erythroid and other myeloid colony replating. The red color of erythroid colonies appears to be more vivid in cultures containing PDS. Premade methylcellulose mixture (Methocult GF M3434, cat. no. 03434), purchased from StemCell Technologies, can also be successfully used for replating day-4 and day-9 EBs.

Keller, G., Kennedy, M., Papayannopoulou, T., and Wiles, M.V. 1993. Hematopoietic commitment during embryonic stem cell differentiation in culture. Mol. Cell. Biol. 13:473-486.

Anticipated Results

We typically analyze FLK1+ cells from day-3 to day-5 EBs. For R1 ES cells, FLK1+ cells represent ∼10% in day-3 EBs; ∼30% to 50% in day-4 EBs; and ∼20% in day 5 EBs. Blast colony forming cells (BL-CFCs) typically represent ∼1% to 3% of day-2.75 to day-3 EBs. Primitive erythroid progenitors represent ∼10% of day-4 EBs. Definitive hematopoietic progenitors represent about 1% of day-6 to day-7 EBs. About 4% to 7% and 2% to 4% of day-6 EBs express CD45 and TER119, respectively (Zhang et al., 2005). It is important to note, however, that the kinetics of FLK1 expression as well as hematopoietic progenitor development can be different among different ES lines. Individual lines need to be examined independently.

Time Considerations Differentiation of mESCs to Blood

We typically set up 2 to 3 consecutive differentiations once ES cells are thawed. It takes about 3 to 4 weeks to complete one round of 2

Literature Cited Choi, K., Kennedy, M., Kazarov, A., Papadimitriou, J., and Keller, G. 1998. A common precursor for hematopoietic and endothelial cells. Development 125:725-732. Chung, Y.S., Zhang, W.J., Arentson, E., Kingsley, P.D., Palis, J., and Choi, K. 2002. Lineage analysis of the hemangioblast as defined by FLK1 and SCL expression. Development 129:5511-5520. Karasuyama, H. and Melchers, F. 1988. Establishment of mouse cell lines which constitutively secrete large quantities of interleukin 2, 3, 4 or 5, using modified cDNA expression vectors. Eur. J. Immunol. 18:97-104.

Kennedy, M., Firpo, M., Choi, K., Wall, C., Robertson, S., Kabrun, N., and Keller, G. 1997. A common precursor for primitive erythropoiesis and definitive haematopoiesis. Nature 386:488-493. Lugus, J.J., Chung, Y.S., Mills, J.C., Kim, S.I., Grass, J., Kyba, M., Doherty, J.M., Bresnick, E.H., and Choi, K. 2007. GATA2 functions at multiple steps in hemangioblast development and differentiation. Development 134:393405. Nakano, T., Kodama, H., and Honjo, T. 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265:10981101. Nishikawa, S.I., Nishikawa, S., Hirashima, M., Matsuyoshi, N., and Kodama, H. 1998. Progressive lineage analysis by cell sorting and culture identifies FLK1+VE-cadherin+ cells at a diverging point of endothelial and hemopoietic lineages. Development 125:1747-1757. Okuda, T., van Deursen, J., Hiebert, S.W., Grosveld, G., and Downing, J.R. 1996. AML1, the target of multiple chromosomal translocations in human leukemia, is essential for normal fetal liver hematopoiesis. Cell 84:321-330. Park, C., Afrikanova, I., Chung, Y.S., Zhang, W.J., Arentson, E., Fong, Gh G., Rosendahl, A., and Choi, K. 2004. A hierarchical order of factors in the generation of FLK1- and SCL-expressing hematopoietic and endothelial progenitors from embryonic stem cells. Development 131:27492762. Park, C., Lugus, J.J., and Choi, K. 2005. Stepwise commitment from embryonic stem to hematopoietic and endothelial cells. Curr. Top. Dev. Biol. 66:1-36. Pevny, L., Simon, M.C., Robertson, E., Klein, W.H., Tsai, S.F., D’Agati, V., Orkin, S.H., and Costantini, F. 1991. Erythroid differentiation in chimaeric mice blocked by a targeted mutation in the gene for transcription factor GATA-1. Nature 349:257-260.

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Qi, X., Li, T.G., Hao, J., Hu, J., Wang, J., Simmons, H., Miura, S., Mishina, Y., and Zhao, G.Q. 2004. BMP4 supports self-renewal of embryonic stem cells by inhibiting mitogen-activated protein kinase pathways. Proc. Natl. Acad. Sci. U.S.A. 101:6027-6032. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. (eds.). 2008. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Shivdasani, R.A., Mayer, E.L., and Orkin, S.H. 1995. Absence of blood formation in mice lacking the T-cell leukaemia oncoprotein tal-1/SCL. Nature 373:432-434. Tsai, F.Y., Keller, G., Kuo, F.C., Weiss, M., Chen, J., Rosenblatt, M., Alt, F.W., and Orkin, S.H. 1994. An early haematopoietic defect in mice

lacking the transcription factor GATA-2. Nature 371:221-226 Wang, Q., Stacy, T., Binder, M., Marin-Padilla, M., Sharpe, A., and Speck, N. 1996. Disruption of the Cbfa2 gene causes necrosis and hemorrhaging in the central nervous system and blocks definitive hematopoiesis. Proc. Natl. Acad. Sci. U.S.A. 93:3444-3449. Wiles, M.V. and Keller, G. 1991. Multiple hematopoietic lineages develop from embryonic stem (ES) cells in culture. Development 111:259-267. Zhang, W.J., Park, C., Arentson, E., and Choi, K. 2005. Modulation of hematopoietic and endothelial cell differentiation from mouse embryonic stem cells by different culture conditions. Blood 105:111-114.

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Endothelial Differentiation of Embryonic Stem Cells

UNIT 1F.5

Alicia A. Blancas,1 Nicholas E. Lauer,2 and Kara E. McCloskey1, 2 1

Graduate Program in Quantitative and Systems Biology, University of California at Merced, Merced, California 2 School of Engineering, University of California at Merced, Merced, California

ABSTRACT Vascular progenitor cells derived from stem cells could potentially lead to a variety of clinically relevant applications, including cell-based therapies and tissue engineering. Here, we describe methods for isolating purified proliferating populations of vascular endothelial cells from mouse embryonic stem cells (mESC) using Flk-1 positive sorted cells, VEGF supplementation, and a rigorous manual selection technique required for endothelial cell purification and expansion. Using this in vitro derivation procedure, it is possible to obtain millions of cells at various stages of differentiation, with the potential C 2008 for up to 25 population doublings. Curr. Protoc. Stem Cell Biol. 6:1F.5.1-1F.5.19.  by John Wiley & Sons, Inc. Keywords: embryonic stem cells r endothelial cells r endothelial progenitor cells r vascular progenitor cells r Flk-1 r VEGF

INTRODUCTION Vascular endothelial cells or endothelial progenitor cells derived from stem cells could potentially lead to a variety of clinically relevant applications (Dzau et al., 2005). These cells could be used in therapeutic strategies for the repair and revascularization of ischemic tissue in patients exhibiting vascular defects (Kalka et al., 2000; Soker et al., 2000). Endothelial progenitor cell transplantation has been shown to induce new vessel formation in ischemic myocardium and hind limb (Kalka et al., 2000; Kawamoto et al., 2001; Kocher et al., 2001). Since it is well known that endothelial cells inhibit platelet adhesion and clotting, they are needed for lining the lumen of a synthetic or tissue-engineered vascular graft or for re-endothelization of injured vessels (Kaushal et al., 2001; Griese et al., 2003). Moreover, because endothelial cells line the lumen of blood vessels and can release proteins directly into the blood stream, they are ideal candidates to be used as vehicles of gene therapy. Endothelial cells may also be used for vascularizing tissue-engineered materials prior to implantation and for investigating mechanisms of angiogenesis and vasculogenesis. One potential source for these therapeutic endothelial cells is the embryonic stem cell (ESC). The ESC possesses some advantages over adult stem cells in that the ESC provides an excellent in vitro culture system for studying cellular differentiation events, and because the ESC is thought to have the capacity for an unlimited number of cell divisions, it may retain greater potential for in vitro expansion of large numbers of tissue-specific cells. The methodology presented in this unit expands on the work of Nishikawa’s group (Nishikawa et al., 1998, 2001a; Yamashita et al., 2000) for the in vitro differentiation and purification (>96% pure) of EC populations from mouse ESC (McCloskey et al., 2003). These ESC-derived endothelial cells display characteristics of the vascular endothelial cell in that they express several endothelial markers (McCloskey et al., 2003), and they form two-dimensional tube-like structures, as well as complex vessel-like structures in three-dimensional collagen type I gels (McCloskey et al., 2005). Current Protocols in Stem Cell Biology 1F.5.1-1F.5.19 Published online September 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f05s6 C 2008 John Wiley & Sons, Inc. Copyright 

Embryonic and Extraembryonic Stem Cells

1F.5.1 Supplement 6

In this unit, we describe detailed protocols for derivation of EC from mouse ESC (for both R1 and D3 cell lines). We also provide a second protocol for the maintenance and initial differentiation of EC from mouse ESC under serum-free conditions. Lastly, we review current methods of EC differentiation from human ESC. BASIC PROTOCOL

ENDOTHELIAL CELL DIFFERENTIATION FROM MOUSE ESC This first section presents detailed methods for isolating purified proliferating populations of endothelial cells from mouse embryonic stem cells using a 2-D induction on collagen IV, followed by sorting of the Flk-1+ cells that are generated, VEGF supplementation, and a second, more rigorous manual selection technique for isolation of highly purified populations of EC. Using this in vitro derivation procedure, large numbers of endothelial cells can be expanded for up to 25 population doublings. The ESC culturing methods described here provide ∼106 ESC per 35-mm dish at confluence. These small dishes are maintained due to the expense of reagents; however, if larger numbers of cells are desired, this protocol may be scaled up proportionally keeping constant the cell seeding density (the number of cells per cm2 ). Generalized protocols for freezing, thawing, and mitomycin inactivation of cells used in these experiments—feeder cells, ESC, EC—are provided in Support Protocols 1, 2, and 3.

Materials ES-D3 or ES-R1 cells (American Type Culture Collection, cat. no. CRL-1934 or SCRC-1036) Mouse ESC medium (see recipe) Dulbecco’s phosphate-buffered saline (D-PBS), calcium- and magnesium-free (Invitrogen, cat. no. 14190-144) Trypsin/EDTA (Invitrogen, cat. no. 25300-054) ESC-to-EC differentiating medium (see recipe) Gelatin (for subculturing of cells) Cell dissociation solution (Sigma, cat. no. C-5914) Fetal bovine serum (FBS), heat inactivated (Cellgro, cat. no. 35-001-CV) BSA buffer solution (see recipe) Normal donkey serum (Research Diagnostics, cat. no. RDI-NSDNKY) Rabbit anti–mouse Flk-1 (Alpha Diagnostic International, cat. no. FLK11-A) Donkey anti–rabbit phycoerythrin (PE)-conjugated (Research Diagnostics, cat. no. RDI-711116152) Recombinant human vascular endothelial growth factor (VEGF165 ; R&D Systems, cat. no. 293-VE) EC medium (see recipe) Collagen IV (Becton-Dickinson; cat. no. 354233) or collagen I (Becton-Dickinson; cat. no. 354236) or fibronectin (Sigma; cat. no. F-1141) or gelatin (Sigma; cat. no. G-1890) for coating flasks for expansion Gelatin (Sigma, cat. no. G-1890)

Endothelial Differentiation of Embryonic Stem Cells

Fibroblast feeder cell–coated 35-mm dishes (Support Protocol 3) 15-ml centrifuge tubes (VWR, cat. no. 21008-103) Benchtop centrifuge Biocoat collagen IV 35-mm culture dishes (Becton-Dickinson, cat. no. 354459) Cell scraper, optional Vortex 5-ml round-bottomed polystyrene FACS tube Fluorescent-activated cell sorter (FACS) 25-, 75-, and 175-cm2 flasks

1F.5.2 Supplement 6

Current Protocols in Stem Cell Biology

Inverted microscope (for general viewing of cells) Stereomicroscope Additional reagents and equipment for thawing ES-D3 cells (Support Protocol 2), performing a viable cell count (UNIT 1C.3), preparing dissecting pipets (Support Protocol 4), and preparing a mouth aspirator (Support Protocol 5) NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Culture ESC 1. Thaw ES-D3 cells (Support Protocol 2) making sure to use mouse ESC medium. 2. Plate 1 × 105 to 5 × 105 ES-D3 cells per fibroblast feeder cell–coated 35-mm dish with 2.5 ml mouse ESC medium. 3. Replace culture medium daily.

Subculture ESC 4. Subculture the cells before colonies begin to touch. If 2 × 105 cells per 35-mm dish are plated, they will need to be subcultured in 3 days. ES cells maintain their undifferentiated state best when the colonies are subcultured before the colonies come in contact with other colonies (Fig. 1F.5.1).

Figure 1F.5.1 Mouse ESC colonies on embryonic fibroblast feeder cells. Note that, in general, the ESC colonies are not in contact with one another and should be subcultured well before colonies begin to contact one another. This figure shows what is considered a “confluent” dish. These cells should be subcultured within 24 hr. Embryonic and Extraembryonic Stem Cells

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5. Gently remove culture medium and rinse cells twice, each time with 3 ml D-PBS per 35-mm dish. 6. Add 1 ml trypsin/EDTA per 35-mm dish and place cells in the incubator for 1 to 2 min. 7. Gently pipet cells up and down 10 times to disaggregate cells. 8. Add 3 ml mouse ESC medium and transfer all of the cell suspension to a 15-ml centrifuge tube. 9. Add an additional 3 to 5 ml mouse ESC medium to completely neutralize the trypsin. 10. Again, gently pipet cells up and down an additional 10 to 30 times to disaggregate cells. The goal is to obtain a single-cell suspension. Because fibroblast feeders tend to stick together, allow 2 min for the large cell clumps to sink to the bottom of the centrifuge tube. Then, transfer the top 34 of the cell suspension to another centrifuge tube and discard the fibroblast cell clumps in the first tube. This technique also ensures that fewer fibroblasts are subcultured in the next dish.

11. Count cells (UNIT 1C.3). 12. Centrifuge 4 to 5 min at 200 × g, room temperature. 13. Remove supernatant. 14. Resuspend pellet in appropriate quantities of mouse ESC medium and replate at 1 × 105 to 5 × 105 ES-D3 cells per fibroblast feeder cell–coated 35-mm dish with 2.5 ml mouse ESC medium per dish.

Collect ESC 15. Gently remove culture medium and rinse cells twice, each time with 3 ml D-PBS per 35-mm dish. 16. Add 1 ml trypsin/EDTA per 35-mm dish and place cells in the incubator for 1 to 2 min. 17. Gently pipet cells up and down 10 times to disaggregate cells. 18. Add 3 ml ESC-to-EC differentiating medium and transfer all of the cell suspension to a 15-ml centrifuge tube. 19. Add an additional 3 to 5 ml ESC-to-EC differentiating medium to completely neutralize the trypsin. 20. Again, gently pipet cells up and down an additional 10 to 30 times to disaggregate cells. 21. Centrifuge 4 to 5 min at 200 × g, room temperature. 22. Remove supernatant.

Replate cells for differentiation The cells are subcultured on 0.1% gelatin (no feeders) for 3 to 6 days before switching to differentiation conditions. This allows expansion of the embryonic stem cells, while minimizing the number of feeder cells in the culture. Endothelial Differentiation of Embryonic Stem Cells

23. Resuspend pellet in 1 ml ESC-to-EC differentiating medium and gently pipet cells up and down an additional 10 to 30 times to disaggregate cells. 24. Count cells (UNIT 1C.3).

1F.5.4 Supplement 6

Current Protocols in Stem Cell Biology

Figure 1F.5.2 Mouse ESC colonies on gelatin (A). Mouse ES cells after 3 to 4 days of differentiation on collagen type IV (B). Note the distinct changes in morphology between undifferentiated ES cells and differentiated ES cells.

25. Add 2.5 ml ESC-to-EC differentiating medium to each of 2 to 4 biocoat collagen IV 35-mm culture dishes. 26. Add 30,000 cells (calculated volume) to each 35-mm collagen IV–coated dish. 27. Incubate 4 days at 37◦ C and 5% CO2 . Do not change culture medium during these 3 to 4 days.

Collect Flk-1+ vascular progenitor cells After 3 to 4 days of differentiation, the ESCs will consist of a heterogeneous mixture of progenitor cells. When ESCs begin to differentiate, they will lose typical 3-D colony appearance and begin to grow more like monolayer cell cultures (Fig. 1F.5.2). Included in the mixture will be a population of Flk-1 expressing cells that are vascular progenitor cells and blood precursor cells (for discussion see Nishikawa et al., 1998, 2001a; Hirashima et al., 1999; Yamashita et al., 2000; McCloskey et al., 2003). Using flow cytometry, the brightest Flk-1 expressing cells can be isolated from the heterogeneous mixture of cells. 28. Remove culture medium and wash cells twice, each time with 3 ml of D-PBS per 35-mm dish. 29. Add 3 ml of cell dissociation solution to each dish and allow cells to incubate 20 to 30 min at 37◦ C. When staining cells for extracellular surface markers, it is very important to use a non-enzymatic method for removing the cells from the culture dishes; therefore, do not use trypsin when staining cells. Trypsin will degrade the surface markers that you are attempting to stain.

30. Pipet up and down 10 times while washing solution over the bottom of the dish to remove all the cells. If some cells are still adhering to the bottom of the dish, then use a cell scraper to remove the remaining cells. 31. Transfer cells to a 15-ml centrifuge tube and add 3 ml heat-inactivated FBS. 32. At this stage, pool up to five 35-mm dishes of cells for staining and sorting. 33. Centrifuge cells 4 to 5 min at 200 × g, room temperature. The cells are now ready for immunostaining. Take care to keep the cells sterile during the entire staining and sorting procedure. All solutions for staining will be kept at 4◦ C or on ice.

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Stain cells for sorting 34. Chill the BSA buffer solution at 4◦ C or keep buffer solution on ice. 35. Remove supernatant from the cell pellet. 36. Resuspend the entire cell pellet in 1 ml BSA buffer solution/10% donkey serum. Vortex gently. 37. Incubate 1 hr on ice or at 4◦ C. 38. Add 4 ml of BSA buffer solution and centrifuge 4 to 5 min at 200 × g, 4◦ C. Vortex gently. 39. Remove supernatant and resuspend the cell pellet in 400 μl of BSA buffer solution. Pipet up and down to evenly distribute the cells in the solution.

Expose cells to Flk-1 antibody 40. Place 50 μl of cell suspension in another 15-ml centrifuge tube and label it “cells only.” 41. Place 50 μl of cell suspension in another 15-ml centrifuge tube and label it “PE only” (or use an IgG PE isotype control). 42. Label the original cell suspension “Flk-1 PE.” 43. Add 250 μl BSA buffer to the two new centrifuge tubes. All tubes should now be at 300 μl.

44. Add 8 μl of Flk-1 antibody to the tube labeled “Flk-1 PE.” 45. Incubate all tubes 30 min on ice or at 4◦ C. 46. Add 4 ml BSA buffer solution to all tubes and centrifuge 4 to 5 min at 200 × g, 4◦ C. 47. Remove supernatant and resuspend the cell pellets, each in 300 μl of BSA buffer solution. Pipet up and down to evenly distribute the cells in the solution.

Expose cells to secondary antibody 48. Add 8 μl of donkey anti–rabbit PE to the tube labeled “Flk-1 PE” and the tube labeled “PE only.” Alternatively, use an IgG PE isotype control. Fluorescent antibodies should be kept in the dark during storage and when labeling cells. Exposure to too much light may cause the fluorescent molecules to emit light prematurely.

49. Incubate all tubes 30 min on ice or at 4◦ C. 50. Add 4 ml BSA buffer solution to all tubes and centrifuge 4 to 5 min at 200 × g, 4◦ C. 51. Remove supernatant and repeat step 50.

Sort the cells 52. Resuspend the cells in the “Flk-1 PE” tube in 1 ml of BSA buffer and transfer the cell solution to a labeled 5-ml round-bottomed polystyrene FACS tube. 53. Resuspend the cells in “cells only” and “PE only” tubes in 300 μl of BSA buffer and transfer the cell solutions to labeled 5-ml round-bottomed polystyrene FACS tubes. 54. Fill a fourth 5-ml round-bottomed polystyrene FACS tube with 1.5 ml of ESC-to-EC differentiating medium. Endothelial Differentiation of Embryonic Stem Cells

This tube will serve as your “collection” tube for fluorescence-activated cell sorting (FACS).

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Figure 1F.5.3 R1 ESC exhibit high Flk-1 expression after 3 days on collagen type IV (A) or gelatin (B). These are the vascular progenitor cells that will be isolated from the heterogeneous mixture of cells.

55. Sort the “brightest” population of Flk-1 expressing cells into the “collection” tube. Usually there will be a subpopulation of cells that is expressing a very high number of Flk-1 surface molecules. This population will be the “brightest” population of cells falling in the highest channels of your FACS histogram. This population of cells typically ranges from 10% to 30% of your total cell population (Fig. 1F.5.3).

Plate and culture the Flk-1+ cells 56. Centrifuge the “collection” tube containing Flk-1 positive cells 3 to 4 min at 200 × g, room temperature. 57. Remove the supernatant and resuspend cells in 1 ml of ESC-to-EC differentiating medium. Based on number of cell-sorting events, calculate the volume of cell suspension to add to each 35-mm collagen IV-coated dish. You will want ∼50,000 to 100,000 cells per dish.

58. Add 2.5 ml of ESC-to-EC differentiating medium to each dish. 59. Add 125 μl of VEGF (50 ng/ml) to each dish. 60. Put the cells in a 37◦ C incubator and do not move dishes for 4 days. 61. On day 4, aspirate off old medium and add 2.5 ml fresh ESC-to-EC differentiating medium plus 125 μl of VEGF per 35-mm dish. Resume incubation. Most of these cells will die due to staining and FACS sorting procedures. Do not move the dishes or change medium for 4 days and then allow at least 1 week before expecting to see any cell growth.

Purify ECs from vascular progenitor cells After culturing for ∼1 week, the Flk-1 positive cell outgrowths exhibit predominantly two different morphologies (see Fig. 1F.5.4). These include endothelial-like cells with a cobblestone morphology, and elongated smooth muscle-like cell populations. Since these two populations are distinctly different in appearance, it is possible to manually isolate the endothelial cells and replate them in clean dishes for further purification.

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Figure 1F.5.4 Outgrowths of Flk-1 positive cells consist of primarily two cell populations: endothelial-like cells exhibiting a cobblestone-like morphology (A) stained with endothelial marker PE-CAM1 (B), and elongated smooth muscle-like cells (C) stained with alpha-smooth muscle actin (D).

62. Prepare dissecting pipets (Support Protocol 4) and mouth aspirator (Support Protocol 5). 63. Aspirate culture medium and wash cells twice, each time with 3 ml of D-PBS per 35-mm dish. 64. Incubate cells 5 min with cell dissociation solution. As cells begin to detach from the culture dish, their distinct cell morphologies may become vague. It is helpful to mark the bottom of the dish with the appropriate location of the desired cells and work quickly.

65. Meanwhile, fill 6 to 10 collagen IV-coated 35-mm dishes with 2 ml of EC medium. 66. Using a stereomicroscope for optimal visualization, carve around a 5 to 10 cell cluster with the edge of the mouth-Pasteur pipet assembly (see Fig. 1F.5.5).

Endothelial Differentiation of Embryonic Stem Cells

67. Aspirate the cells into the pipet and transfer to the new 35-mm dishes containing 2 ml of ESC-EC differentiation medium. 68. Repeat carving out another 5 to 10 cluster of cells and plate in a new 35-mm dish.

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Figure 1F.5.5 Endothelial-like cells exhibiting a cobblestone-like morphology are manually picked based on proper morphology and replated on a second dish coated with collagen IV (A) and a photograph of the aspiration device used for the manual picking (B). Note that several batches of endothelial cells may be isolated from one dish. These batches may vary slightly, so it is a good idea to expand the batches separately.

69. Repeat for 6 to 10 dishes, using a separate dish for each cluster. The number of clusters obtained depends on the quality of the EC sheets, 6 to 10 clusters per 35-mm dish of Flk-1+ outgrowths is normal.

70. Add 50 ng/ml of VEGF to each dish. 71. Incubate cells 7 to 10 days at 37◦ C and 5% CO2 . Change medium every 4 days.

Expand ECs in vitro 72. After allowing the cells 7 to 10 days of uninterrupted growth, observe the dishes carefully for EC colonies. Once the cell colonies are well established, you will see 50 to 100 cells in a circular sheet. These cells will be highly confluent in the center and appear to grow outward at the edges of the colony.

73. To encourage further cell proliferation, subculture the cells using enzymatic passaging in EC medium to allow cells to grow easily on the entire surface of the collagen IV-coated 35-mm dish. 74. Continually expand the cells in larger dishes (35-mm dish, then 25-cm2 flask, then 75-cm2 flask, then 175-cm2 flask, and then multiple 175-cm2 flasks). Make sure to coat the surface of each flask with collagen IV, collagen I, fibronectin, or gelatin for 2 hr prior to cell seeding and wash off the extra substrate with PBS. The ESC derived EC can be frozen and thawed normally (Support Protocols 1 and 2). Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 1

FREEZING CULTURED CELLS Feeder cells, ESCs, and ECs can be frozen to maintain stocks of cells until they are needed. This is a generalized procedure for freezing cells. The cells are removed from the dish, resuspended in freezing medium and frozen.

Materials Cultures to be frozen Trypsin/EDTA (Invitrogen, cat. no. 25300-054) Phosphate-buffered saline, calcium- and magnesium-free Appropriate medium for cells containing serum Freezing medium (see recipe) 35-mm tissue culture dishes Phase contrast microscope Nunc cryovials (VWR, cat. no. 66021-986) Cryo 1◦ C freezing containers (Research Products International, cat. no. 5100-0001) −70◦ or −80◦ C freezer Liquid nitrogen storage tank 1. Trypsinize cells in the exponential phase of growth (varies for each cell type, but typically is after 3 days of growth). First aspirate the medium and wash the culture twice, each time with 3 ml of PBS per 35-mm dish, and then add 1 ml trypsin/EDTA. Incubate under the phase contrast microscope. After ∼3 min cells begin to round with clearly defined edges.

2. Once cell rounding is observed, add 3 ml of medium with serum and pipet several times to disaggregate cells from the dish and from each other until a single-cell suspension is achieved. This is a general trypsinization procedure. The medium added after trypsinization should be the same as the cells are currently cultured in.

3. Pellet cells by centrifuging 4 to 5 min at 200 × g, room temperature and resuspend in an appropriate amount of cell culture medium. Count cells. For convenience, cells are frozen in 1-ml aliquots at cell numbers that correspond to the appropriate numbers that will be needed upon thawing. The upper limit would be 5 to 10 × 106 cells/ml.

4. Slowly add an equal volume of the freezing medium dropwise over 2 min. Continuously shake the cell suspension for even distribution of the freezing medium. 5. Divide cell suspension into 1-ml aliquots into cryovials. ESC are typically frozen between 5 × 105 and 1 × 106 cells/ml.

6. Immediately transfer cryovials to a cryo 1◦ C freezing container and place the container in a −70◦ C or a −80◦ C freezer for 24 hr. 7. Transfer the vials to liquid nitrogen storage tank. SUPPORT PROTOCOL 2

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THAWING CULTURED CELLS Frozen stocks of cultured cells need to be carefully thawed to ensure viability. This is a generalized method applicable to feeder cells, ESCs, and ECs.

Materials Frozen stocks of cells (Support Protocol 1) Appropriate cell medium

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37◦ C water bath Laminar flow cabinet 15-ml centrifuge tube 1. Thaw the cells in a 37◦ C water bath until only a small ice droplet remains (∼1 min, the last drop will thaw as you carry the vial to the laminar flow cabinet). 2. While the vial is thawing, fill a 15-ml centrifuge tube with 10 ml of the appropriate cell culture medium for the cell types. You will use embryonic fibroblast feeder cell medium for fibroblasts, ESC medium for mouse ESC, and EC medium for fully differentiated and purified EC. Cells at intermediate stages of differentiation are not usually frozen.

3. Transfer thawed cells to the centrifuge tube and collect the cells by centrifuging 4 to 5 min at 200 × g, room temperature. 4. Remove the supernatant and gently resuspend the cells in 4 to 5 ml fresh growth medium. 5. Transfer cells to the prepared culture dish and place in a 37◦ C incubator. ESC should be plated at 1 × 105 cells per 35-mm dish. Fibroblast feeder cells should be plated at 4 × 105 cells per 35-mm dish. Both ESC and EC cells are maintained on dishes or flasks coated with the appropriate substrate; therefore, when thawing or passing cells, make sure to have allowed time (1 to 2 hr) for the substrate to adhere to the culture dish and wash off excess substrate with PBS. For ESCs that will be cultured on fibroblasts, make sure to prepare those dishes with a layer of fibroblast cells at least 4 hr prior to ESC seeding.

6. Replace the medium with fresh ESC medium the next day.

MITOTIC INACTIVATION OF FIBROBLAST FEEDER Typically, ES cells are cultured on fibroblast feeder cells that are inactivated with mitomycin C or irradiation. The inactivation of the fibroblast cells allows the ES cells to benefit from the co-culture feeder conditions without fibroblast proliferation. Mouse embryonic fibroblast feeder cells are typically used; however, the isolation of these cells requires several animals to be sacrificed and labor-intensive dissection of the fetal tissue. If mouse embryonic feeders are unavailable, or undesirable, STO cells may also be used (available from ATCC). Before disposing, mitomycin C must be neutralized with Clorox bleach for at least 15 min.

SUPPORT PROTOCOL 3

Materials Feeder cells to be inactivated: mouse fibroblasts or STO cells (ATCC, cat. no. CRL-1503) Embryonic fibroblast feeder cell medium (see recipe) Mitomycin C solution (see recipe) Phosphate-buffered saline (PBS), with calcium and magnesium Phosphate-buffered saline (PBS), calcium- and magnesium free Trypsin/EDTA 175-cm2 tissue culture flasks (with 0.2-mm vent cap; Corning, cat. no. 431080) 37◦ C incubator 15-ml centrifuge tubes 35-mm dish Additional reagents and equipment for counting cells (UNIT 1C.3)

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Grow fibroblasts 1. After thawing (Support Protocol 2), allow mouse fibroblasts to grow to 90% to 95% confluency in 175-cm2 tissue-culture treated flasks in embryonic fibroblast feeder cell medium. Inactivate cells 2. Aspirate culture medium from flask and replace with 16 ml of mitomycin C solution. 3. Incubate the treated flasks 2 hr at 37◦ C, 5% CO2 . 4. After 2 hr, aspirate mitomycin C solution and wash each 175-cm2 flask five times, four times with 20 ml PBS with calcium and magnesium and once (last wash) with calcium- and magnesium-free PBS. 5. Add 3 ml trypsin/EDTA per flask and monitor cell detachment. After ∼1 min, cells should detach from the flask surface (gently rock flask side-to-side).

6. After cells have detached, add 5 to 10 ml of embryonic fibroblast feeder cell medium. 7. Transfer the cell suspension from each flask to 15-ml centrifuge tubes. 8. Centrifuge 4 to 5 min at 200 × g, room temperature. 9. Remove supernatant and wash again with 10 ml embryonic fibroblast feeder cell medium per tube. 10. Centrifuge 4 to 5 min at 200 × g, room temperature. 11. Repeat washing one more time. Resuspend the cells in 1 ml embryonic fibroblast feeder cell medium. 12. Count cells (UNIT 1C.3).

Plate cells 13. Plate between 3 × 105 and 4 × 105 cells per 35-mm dish that will be needed for ESC culture. Add 3 ml embryonic fibroblast feeder cell medium to each dish and allow at least 4 hr, preferably overnight, for the cells to adhere to dishes before adding embryonic stem cells. Excess inactivated fibroblasts may also be frozen at this point for future use. Inactivated fibroblasts may be used for up to 1 week. SUPPORT PROTOCOL 4

PREPARATION OF DISSECTING PIPETS Pipets must be modified for manual dissection of EC progenitor cells for passaging.

Materials Glass Pasteur pipets (9 in.; VWR, cat. no. 53283-915) Bunsen burner 1. Hold the narrow tip of a Pasteur pipet in your left hand and larger end in your right hand. Pass the center of the narrow portion through a low flame of a Bunsen burner until the pipet is hot. 2. Quickly pull on the tip of the pipet while lifting the pipet out of the flame to generate a pipet region with a smaller diameter just above the tip of the pipet. Endothelial Differentiation of Embryonic Stem Cells

3. Loop back the pulled glass and rub glass to glass to create a point of friction. Tap the glass to break the tip off at the point of friction. The technique for pulling Pasteur pipets will take some practice.

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4. Polish the new end of the pipet by passing the new tip gently over a low flame. The Pasteur pipets should remain sterile, so use immediately after pulling, or pull several pipets and sterilize them ahead of time.

PREPARING A MOUTH ASPIRATOR A mouth aspirator is used with the dissecting pipet when passaging EC progenitor cells.

SUPPORT PROTOCOL 5

Materials 1000-μl micropipet tip Aspirator assembly with rubber tubing (Sigma, cat. no. A5177) 0.2-μm syringe filter (Pall, cat. no. 4192) Dissecting pipet (Support Protocol 4) 1. Fit the narrow end of a 1000-μl micropipet tip into the rubbing tubing of an aspirator assembly fitted with a 0.2-μm syringe filter. 2. Insert the modified Pasteur dissecting pipet into the wide end of the 1000-μl micropipet tip. This aspirator assembly allows for simultaneous microscope viewing and cell colony manipulations.

EC DIFFERENTIATION FROM MOUSE ESC CULTURE UNDER SERUM-FREE CONDITIONS

ALTERNATE PROTOCOL 1

The methods described above employ methods of cell culture and differentiation where the ESC are grown in medium containing fetal bovine serum (FBS). However, the reproducibility of some aspects of these experiments can vary since FBS composition can vary significantly from batch-to-batch. This leads to tiresome batch testing and buying up entire lots of screened batches of FBS at once. This process must then be repeated when the desired lot is exhausted. By using an induction system that does not require serum, the conditions under which the cells are grown are chemically defined, and more reproducible. Based on the formulas previously developed (Adelman et al., 2002; Tanaka et al., 2006), it is possible to maintain murine ESC in culture on gelatin in a chemically defined serum-free medium. The cells retain their morphology well and replicate quickly with a doubling time of ∼3 days (this is a slower growth rate than achieved with serum). Efforts to develop a chemically defined medium for differentiation have been more difficult. In the absence of serum and LIF, the cells differentiate, but proliferate much more slowly in comparison to the induction medium with serum. However, the percentage of Flk-1+ cells in the serum-free induction is comparable to that obtained from inductions with serum-containing medium and, therefore, can be scaled-up to achieve the desired number of Flk-1+ cells. EC differentiation in a two-dimensional system has been traditionally performed on collagen IV-coated dishes on the premise that collagen IV induces the greatest number of mesodermal cells (Nishikawa et al., 2001b, 2007). However, our laboratory has succeeded in inducing equally sufficient expression of Flk-1 on gelatin-coated dishes. When compared to a serum-containing differentiation medium, our serum-free mixture yielded a comparable percentage of Flk-1+ cells, 20%. Embryonic and Extraembryonic Stem Cells

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Additional Materials (also see Basic Protocol) Bone morphogenic protein 4 (BMP-4; R&D Systems) Serum-free ESC culture medium (see recipe) Serum-free ESC differentiating medium (see recipe) The basic steps for serum-free culture and EC induction follow those of the Basic Protocol. Substitute the serum-free ESC culture medium in the steps for ESC culture (steps 1 to 22). Substitute the serum-free ESC differentiating medium in the steps for differentiating ESC and purification of ECs (steps 23 to 71). Serum-free subculture for mature EC (steps 72 to 74) is currently under investigation.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

BSA buffer solution Add 0.6 g bovine albumin (Sigma, cat. no. A-1470) to 200 ml calcium- and magnesium-free PBS to make 0.3% BSA buffer solution. Place this mixture in the 37◦ C water bath until the albumin is dissolved (sterile filter the solution if needed). Store up to 1 week at 4◦ C.

EC medium This is a commercially available EC medium kit; EGM-2 medium Bullet Kit (500-ml bottle plus growth factors; Clonetics, cat. no. CC-3162).

Embryonic fibroblast feeder cell medium 88% (v/v) high-glucose Dulbecco’s modified eagle medium (DMEM; Invitrogen, cat. no. 119650-092) 10% (v/v) heat-inactivated fetal bovine serum (FBS; Cellgro, cat. no. 35-001-CV) 1% penicillin/streptomycin (from 100× stock; Invitrogen, cat. no. 15070-063) 1% (v/v) L-glutamine (from 100× stock; Invitrogen, cat. no. 25030-081) Store for up to 1 month at 4◦ C ESC-to-EC differentiating medium 93% (v/v) α-minimal essential medium (Invitrogen, cat. no.12561-056) 5% (v/v) heat-inactivated fetal bovine serum (FBS; Cellgro, cat. no. 35-001-CV) 1% (v/v) penicillin/streptomycin (from 100× stock; Invitrogen, cat. no. 15070-063) 1% (v/v) L-glutamine (from 100× stock; Invitrogen, cat. no. 25030-081) 5 × 10−5 M 2-mercaptoethanol (Sigma, cat. no. M-7522) Store for up to 1 month at 4◦ C Freezing medium 80% (v/v) heat-inactivated fetal bovine serum (FBS; Cellgro, cat. no. 35-001-CV) 20% (v/v) dimethyl sulfoxide (DMSO; Sigma, cat. no. D2650) Prepare fresh prior to each use To freeze cells, mix equal volumes of the appropriate cell culture medium and freezing medium.

Mitomycin-C solution Endothelial Differentiation of Embryonic Stem Cells

Dissolve 2.0 mg of mitomycin C powder (Sigma, cat. no. M4287) in 200 ml of embryonic fibroblast feeder cell medium (10 μg/ml; see recipe). Stock may be stored up to 6 weeks in the dark at 4◦ C, or at −20◦ C for long-term storage.

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Mouse ESC medium 78% (v/v) Knockout Dulbecco’s modified eagle medium (KO-DMEM; Invitrogen, cat. no. 10829-018) 15% (v/v) ES cell-qualified fetal bovine serum (Invitrogen, cat. no. 16141-079) 5% (v/v) Knockout serum replacement (KSR; Invitrogen, cat. no. 10828-028) 1% (v/v) penicillin/streptomycin (from 100× stock; Invitrogen, cat. no. 15070-063) 1% (v/v) L-glutamine (from 100× stock; Invitrogen, cat. no. 25030-081) 1000 U/ml leukemia inhibitory factor (LIF; Chemicon International, cat. no. ESG1106) 5 × 10−5 M 2-mercaptoethanol (Sigma, cat. no. M-7522) Store for up to 2 weeks at 4◦ C. Serum-free ESC culture medium 15% (v/v) Knockout serum replacement (KSR; Invitrogen, cat. no. 10828-028) 1× penicillin-streptomycin (from 100× stock; Invitrogen, cat. no. 15070-063) 2 mM L-glutamine (from 100× stock; Invitrogen, cat. no. 25030-081) 1× MEM non-essential amino acids (from a 100 × stock from Invitrogen, cat. no. 11140-050) 0.1 mM 2-mercaptoethanol (Calbiochem) 2000 U/ml leukemia inhibitory factor (LIF; Chemicon International, cat. no. ESG1106) 10 ng/ml bone morphogenic protein 4 (BMP-4; R&D Systems) Knockout Dulbecco’s modified eagle medium (KO-DMEM; Invitrogen, cat. no. 10829-018) Serum-free ESC differentiating medium 20% (v/v) Knockout serum replacement (KRS; Invitrogen, cat. no. 10828-028) 1× penicillin/streptomycin (from 100× stock; Invitrogen, cat. no. 15070-063) 2 mM L-glutamine (from 100× stock; Invitrogen, cat. no. 25030-081) 1× MEM non-essential amino acids (from a 100 × stock from Invitrogen, cat. no. 11140-050) 5 × 10−5 M 2-mercaptoethanol (Calbiochem) 5 ng/ml bone morphogenic protein 4 (BMP-4; R&D Systems) 30 ng/ml vascular endothelial growth factor (VEGF; R&D Systems, cat. no. 293VE) α-minimum essential medium (α-MEM; Cellgro) A chemically defined medium (CDM) has also been used for serum-free induction (Johansson and Wiles, 1995; Wiles and Johansson, 1999; Ng et al., 2005). In the cited study, the addition of BMP-4 or Activin A was found to enhance mesoderm differentiation (Johansson and Wiles, 1995).

COMMENTARY Background Information Endothelial cells have been derived from mouse and human ESC by isolating the differentiating endothelium from an embryoid body (Levenberg et al., 2002). Although the embryoid body system enables investigation of vasculogenesis virtually as it occurs in the embryo (Risau et al., 1988; Wang et al., 1992; Vittet et al., 1996; Choi et al., 1998), the multiple cell-cell contacts and cell lineages make it difficult to study and control the behavior of the maturing endothelial cell in detail.

Endothelial, hematopoietic, and smooth muscle cells have also been derived from Flk1+ outgrowths from murine ESCs grown on type-IV collagen-coated surfaces (Nishikawa et al., 1998; Yamashita et al., 2000; McCloskey et al., 2003), showing that the threedimensional structure is not necessary for endothelial maturation from ESC (Nishikawa et al., 1998). The two-dimensional monolayer technique of endothelial differentiation not only allows closer study and control of the in vitro maturation, molecular events, and

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growth factor requirements of endothelial cell derivation (Nishikawa et al., 1998; Hirashima et al., 1999; Yamashita et al., 2000), but also uses an induction method that is devoid of the three-dimensional embryo-like self-programmed machinery for vascular differentiation. Although the two-dimensional monolayer derivation methods have been very successful in isolating and studying the maturation of endothelial cells from murine ESCs, the long-term maintenance of these murine ESC-derived endothelial cells has been limited. Without genetic manipulation, the longest these ESC-derived EC were maintained in culture was 7 days, increasing to two or three passages by culturing cells on OP9 stromal cells (Nishikawa et al., 2001a). In addition to the limitations in the proliferative capabilities of the endothelial cells from murine ESCs (Nishikawa et al., 1998; Hirashima et al., 1999; Yamashita et al., 2000), the reported studies did not isolate uniform populations of endothelial cells from the contaminating smooth muscle cell, or other cell populations. Based on our studies (techniques presented in this unit), the isolation of pure populations of EC is critical for further expansion of these cells (McCloskey et al., 2003). Pure cell populations are also essential for studying the effectiveness of these cells for cell-based therapies and should alleviate the problem of teratomas that form when ESC are implanted in vivo. Recent discoveries of molecular markers for arterial, venous, and lymphatic endothelial cells allow a more sophisticated characterization of endothelial diversity (Aranguren et al., 2007; Yamashita, 2007). Arterial specification, promoted by Notch signaling, is characterized by ephrinB2, Delta-like (Dll)-4, Notch-1 and 4, Jagged-1, and connexin-40 expression. Venous endothelium, potentially a default pathway of EC differentiation, is characterized by EphB4 and COUP-TFII. Committed lymphatic EC, differentiated from venous EC, express Prox-1 as the most specific lymphatic EC marker.

bryonic antigens (SSEA) also varies. Undifferentiated human ESC express SSEA-3 and -4, and do not express SSEA-1, while mouse ESC express SSEA-1 and do not express SSEA-3 or -4. Most importantly, for hESC culture, the presence of LIF does not support undifferentiated feeder-free growth, while LIF is sufficient in mouse ESC cultures. EC differentiation and isolation from hESC was first published by the Langer laboratory in 2002 (Levenberg et al., 2002). In this study, embryoid bodies (EBs) were employed for the initial induction of EC. Endothelial markers CD31, CD34, and VE-cadherin peaked between days 13 and 15 of induction. Sorting CD31+ cells on day 13 allowed for expansion of EC progenitors. After several passages in culture, 78% of the cells still expressed CD31. More recently, the same laboratory used similar protocols for generation of EBs, but sorted CD34+ cells on day 10 to generate vascular progenitor cells retaining the potential to generate both endothelial and smooth muscle cells (Ferreira et al., 2007). Because the formation of EBs from ESC triggers spontaneous differentiation of all cell types, it is an inefficient method for the generation of specific cell types because the microenvironment within the EB is difficult to control. Methods for two-dimensional induction of hESC to EC have also been published (Wang et al., 2007). In this study, hESC were placed on mouse embryonic feeders in differentiation medium containing 15% fetal bovine serum (FBS) for 10 days. By day 10, 5% to 10% of these cells expressed CD34, a common hematopoietic and endothelial progenitor marker. Two rounds of magnetic bead sorting enriched the cells to 80% to 95% purity. When these cells were cultured in endothelial growth medium, the majority of the cells expressed endothelial markers CD31 and VE-cadherin. These researchers were also able to remove FBS by substitution with BIT 9500, VEGF, and BMP-4 growth factors for a serum-free induction, and reported a similar number of CD34+ cells.

EC differentiation from human ESC (hESC) Although both mouse and hESC exhibit similar expression of key transcription factors, including Oct-3/4, Nanog, and Sox2, there are some fundamental differences between mouse and human ESC. For example, the population doubling time of hESC is 36 hr compared with 12 hr for mouse ESC. The hESC grow in relatively flat compact colonies compared with mouse ESC. Expression of stage-specific em-

Critical Parameters The optimal day of initial induction of Flk-1+ vascular progenitor cells is a very small window (<1 day) and will depend on the cell line and culture conditions. ESC-D3 cells will take ∼4 days for optimal induction, while ESC-R1 cells will take only 3 days to generate a significant number of very bright (high expressing) Flk-1+ vascular progenitors. Past this optimal window, the Flk-1+ expression is

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reduced and most of the vascular progenitor cells will be lost. This stage is very important and should be optimized (induce using the ESC induction medium for 2, 2.5, 3, 3.5, 4, and 4.5 days) for the cells before attempting to move forward with the cell sorting of Flk-1+ cells.

Anticipated Results The methods presented here can generate up to 1 to 3 billion EC from each initial induction of 30,000 mouse ESC.

Time Considerations The differentiation, isolation, and expansion procedures (Fig. 1F.5.6) for the generation

Figure 1F.5.6

of EC from mouse ESC will take ∼1 month, but new practitioners should allow another few months for optimization of the methods described. Allow at least 2 weeks, or more, for optimization of the timing for the generation of Flk-1+ cells and sorting these cells. Add another month for practicing the manual selection technique described. Fluorescenceactivated cell sorting or magnetic cell sorting can also be used for selection of EC progenitors from smooth muscle cells, but will not yield the high purity needed for a significant expansion of EC. If the two cell types are not purified away from each other, smooth muscle cells will eventually outgrow EC and the cell cultures will need to be discarded.

Flow diagram of the steps for EC differentiation from mouse ESC. Embryonic and Extraembryonic Stem Cells

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Literature Cited Adelman, C.A., Chattopadhyay, S., and Bieker, J.J. 2002. The BMP/BMPR/Smad pathway directs expression of the erythroid-specific EKLF and GATA1 transcription factors during embryoid body differentiation in serum-free media. Development 129:539-549. Aranguren, X.L., Luttun, A., Clavel, C., Moreno, C., Abizanda, G., Barajas, M.A., Pelacho, B., Uriz, M., Arana, M., Echavarri, A., Soriano, M., Andreu, E.J., Merino, J., Garcia-Verdugo, J.M., Verfaillie, C.M., and Prosper, E. 2007. In vitro and in vivo arterial differentiation of human multipotent adult progenitor cells. Blood 109:26342642.

Kocher, A.A., Schuster, M.D., Szabolcs, M.J., Takuma, S., Burkhoff, D., Wang, J., Homma, S., Edwards, N.M., and Itescu, S. 2001. Neovascularization of ischemic myocardium by human bone-marrow-derived angioblasts prevents cardiomyocyte apoptosis, reduces remodeling and improves cardiac function. Nat. Med. 7:430436. Levenberg, S., Golub, J.S., Amit, M., ItskovitzEldor, J., and Langer, R. 2002. Endothelial cells derived from human embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 99:4391-4396.

Choi, K, Kennedy, M., Kazarov, A., Papadimitriou, J.C., and Keller, G. 1998. A common precursor for hematopoietic and endothelial cells. Development 125:725-32.

McCloskey, K.E., Lyons, I., Rao, R.R., Stice, S.L., and Nerem, R.M. 2003. Purified and proliferating endothelial cells derived and expanded in vitro from embryonic stem cells. Endothelium 10:329-336.

Dzau, V.J., Gnecchi, M., Pachori, A.S., Morello, F., and Melo, L.G. 2005. Therapeutic potential of endothelial progenitor cells in cardiovascular diseases. Hypertension 46:7-18.

McCloskey, K.E., Gilroy, M.E., and Nerem, R.M. 2005. Use of embryonic stem cell-derived endothelial cells as a cell source to generate vessel structures in vitro. Tissue Eng. 11:497-505.

Ferreira, L.S., Gerecht, S., Shieh, H.F., Watson, N., Rupnick, M.A., Dallabrida, S.M., VunjakNovakovic, G., and Langer, R. 2007. Vascular progenitor cells isolated from human embryonic stem cells give rise to endothelial and smooth muscle like cells and form vascular networks in vivo. Circ. Res. 101:286-294.

Ng, E.S., Azzola, L., Sourris, K., Robb, L., Stanley, E.G., and Elefanty, A.G. 2005. The primitive streak gene Mixl1 is required for efficient haematopoiesis and BMP4-induced ventral mesoderm patterning in differentiating ES cells. Development 132:873-884.

Griese, D.P., Ehsan, A., Melo, L.G., Kong, D., Zhang, L., Mann, M.J., Pratt, R.E., Mulligan, R.C., and Dzau, V.J. 2003. Isolation and transplantation of autologous circulating endothelial cells into denuded vessels and prosthetic grafts: Implications for cell-based vascular therapy. Circulation 108:2710-2715. Hirashima, M., Kataoka, H., Nishikawa, S., and Matsuyoshi, N. 1999. Maturation of embryonic stem cells into endothelial cells in an in vitro model of vasculogenesis. Blood 93:1253-1263. Johansson, B.M. and Wiles, M.V. 1995. Evidence for involvement of activin A and bone morphogenetic protein 4 in mammalian mesoderm and hematopoietic development. Mol. Cell Biol. 15:141-151. Kalka, C., Masuda, H., Takahashi, T., Kalka-Moll, W.M., Silver, M., Kearney, M., Li, T., Isner, J.M., and Asahara, T. 2000. Transplantation of ex vivo expanded endothelial progenitor cells for therapeutic neovascularization. Proc. Natl. Acad. Sci. U.S.A. 97:3422-3427. Kaushal, S., Amiel, G.E., Guleserian, K.J., Shapira, O.M., Perry, T., Sutherland, F.W., Rabkin, E., Moran, A.M., Schoen, F.J., Atala, A., Soker, S., Bischoff, J., and Mayer, J.E. Jr. 2001. Functional small-diameter neovessels created using endothelial progenitor cells expanded ex vivo. Nat. Med. 7:1035-1040.

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for myocardial ischemia. Circulation 103:634637.

Kawamoto, A., Gwon, H.C., Iwaguro, H., Yamaguchi, J.I., Uchida, S., Masuda, H., Silver, M., Ma, H., Kearney, M., Isner, J.M., and Asahara, T. 2001. Therapeutic potential of ex vivo expanded endothelial progenitor cells

Nishikawa, S.I., Nishikawa, S., Hirashima, M., Matsuyoshi, N., and Kodama, H. 1998. Progressive lineage analysis by cell sorting and culture identifies FLK1+VE-cadherin+ cells at a diverging point of endothelial and hemopoietic lineages. Development 125:1747-1757. Nishikawa, S.I., Hirashima, M., Nishikawa, S., and Ogawa, M. 2001a. Cell biology of vascular endothelial cells. Ann. N.Y. Acad. Sci. 947:3540. Nishikawa, S.I., Hirashima, M., Nishikawa, S., and Ogawa, M. 2001b. Cell Biology of Vascular Endothelial Cells. Ann. N.Y. Acad. Sci. 947:35. Nishikawa, S.I., Jakt, L.M., and Era, T. 2007. Opinion: Embryonic stem-cell culture as a tool for developmental cell biology. Nat. Rev. Molec. Cell Biol. 8:502-507. Risau, W., Sariola, H., Zerwes, H.G., Sasse, J., Ekblom, P., Kemler, R., and Doetschman, T. 1988. Vasculogenesis and angiogenesis in embryonic-stem-cell-derived embryoid bodies. Development 102:471-478. Soker, S., Machado, M., and Atala, A. 2000. Systems for therapeutic angiogenesis in tissue engineering. World J. Urol. 18:10-18. Tanaka, N., Takeuchi, T., Neri, Q.V., Sills, E.S., and Palermo, G.D. 2006. Laser-assisted blastocyst dissection and subsequent cultivation of embryonic stem cells in a serum/cell free culture system: Applications and preliminary results in a murine model. J. Translat. Med. 4:20. Vittet, D., Prandini, M.H., Berthier, R., Schweitzer, A., Martin-Sisteron, H., Uzan, G., and Dejana, E. 1996. Embryonic stem cells differentiate

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in vitro to endothelial cells through successive maturation steps. Blood 88:3424-3431. Wang, R., Clark, R., and Bautch, V.L. 1992. Embryonic stem cell-derived cystic embryoid bodies form vascular channels: An in vitro model of blood vessel development. Development 114:303-316. Wang, Z.Z., Au, P., Chen, T., Shao, Y., Daheron, L.M., Bai, H., Arzigian, M., Fukumura, D., Jain, R.K., and Scadden, D.T. 2007. Endothelial cells derived from human embryonic stem cells form durable blood vessels in vivo. Nat. Biotechnol. 25:317-318.

Wiles, M.V. and Johansson, B.M. 1999. Embryonic stem cell development in a chemically defined medium. Exp. Cell Res. 247:241-248. Yamashita, J., Itoh, H., Hirashima, M., Ogawa, M., Nishikawa, S., Yurugi, T., Naito, M., and Nakao, K. 2000. Flk1-positive cells derived from embryonic stem cells serve as vascular progenitors. Nature 408:92-96. Yamashita, J.K. 2007. Differentiation of arterial, venous, and lymphatic endothelial cells from vascular progenitors. Trends Cardiovasc. Med. 17:59-63.

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Hematopoietic Differentiation of Human Embryonic Stem Cells by Cocultivation with Stromal Layers

UNIT 1F.6

Katherine L. Hill1 and Dan S. Kaufman1 1

Stem Cell Institute and Department of Medicine, University of Minnesota, Minneapolis, Minnesota

ABSTRACT Human embryonic stem (hES) cells are of remarkable interest both for the utility of these cells for studying basic human developmental biology and as a potential source for novel therapeutics. Here, we provide detailed methodologies of one of the first systems used to mediate differentiation of hES cells—stromal cell coculture. Use of stromal cells adds the ability to manipulate aspects of the developmental niche that support differentiation into a defined lineage. These methods will allow efficient and reproducible development of hematopoietic progenitor cells, as well as potentially mature hematopoietic cells that are suitable for subsequent in vitro and in vivo studies. Curr. Protoc. Stem Cell Biol. C 2008 by John Wiley & Sons, Inc. 6:1F.6.1-1F.6.12.  Keywords: human embryonic stem cells r hematopoietic r stromal r cocultivation

INTRODUCTION Human embryonic stem (hES) cells provide a valuable tool to study human development, produce novel cell and tissue-based therapies, and investigate drug design and safety. In culture, undifferentiated hES cells are capable of long-term self-renewal, as well as maintaining pluripotency (Thomson et al., 1998; Keller, 2005). A major area of interest lies in hematopoietic lineages due to their diverse clinical applications. Indeed, hematopoietic cells are the leading area for cell-based therapies, ranging from hematopoietic stem cells (HSCs) routinely used to assist treatment of malignancies, to red blood cells and platelets used for transfusion medicine therapies, and T lymphocytes and natural killer (NK) cells used for novel immunotherapies. However, there are drawbacks to current sources of all these cells, such as the need for HLA typing, the risk of transmission of infectious disease, etc. Therefore, there is considerable interest in potentially using hES cells as an alternative starting point to derive these hematopoietic lineages. Two approaches are most commonly utilized to differentiate hES cells: embryoid body (EB) formation and stromal cell coculture systems (Kaufman et al., 2001; Chadwick et al., 2003; Vodyanik et al., 2005; Kennedy et al., 2007; Tian and Kaufman, 2008). While each system has advantages and disadvantages, one benefit of differentiating hES cells using stromal cell lines in coculture is the ability to control specific elements that represent components of a micro-environment or anatomical niche that provide support for lineage-specific development. Not only can a specific stromal cell line be selected based upon desired progenitor type, but selected genes can also be up-regulated to further enhance and increase progenitor quality and number. Indeed, in addition to stromal cells that support hematopoiesis (emphasized in this unit), other studies describe different stromal cell lines that support neural and cardiomyocyte development (Mummery et al., 2003, 2007; Zeng et al., 2004). Additionally, stromal cell lines that over-express specific genes, such as Wnts or Notch-ligands have been useful for studies of hematopoietic development (Schmitt et al., 2004; Woll et al., 2008). The relative ease with which these stromal cells can be cultured provides a simple, yet effective method for differentiation Current Protocols in Stem Cell Biology 1F.6.1-1F.6.12 Published online September 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f06s6 C 2008 John Wiley & Sons, Inc. Copyright 

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of hES cells. Coculture experiments are most commonly run for a reasonably short period of time (7 to 21 days). However, due to the nature of in vitro cell culture, some variability in differentiation outcome is inevitable. For example, variation in yield of specific cell populations can occur with different stromal cell passages, plating density of both stromal and hES cells, and number of days the stromal cells are in culture post inactivation. Given these variables, some optimization of this protocol by individual laboratories may be required, based on the cell systems in use and desired progenitor or mature cell type. This protocol describes the use of M2-10B4 (M210) stromal cells as an example of coculture between stromal cell lines and hES cells, and isolation of hematopoietic progenitors through selection of CD34-expressing cells developed in coculture. However, it is known that other stromal cell lines can be cultured in a similar manner and could also be successfully implemented in differentiation studies as well. Indeed, S17 cells and OP9 cells have also been routinely used for derivation of hematopoietic cells from hES cells, and we have previously compared these cells lines with roughly similar effect. However, M210 cells may have a couple of advantages: (1) M210 cells are commercially available through the American Type Culture Collection (ATCC), providing easy access to low-passage cells; and (2) we have recently found that M210 cells are more stable in long-term culture for maintenance of hematopoietic differentiation potential compared to S17 and OP9 cells (K.L.H. and D.S.K., unpub. observ.). hES cell culture is briefly discussed, but additional resources for successful maintenance of undifferentiated hES cells are listed for use as needed. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

COCULTURE OF M210 STROMAL CELLS AND HUMAN ES CELLS Figure 1F.6.1 provides an overview of coculture set up.

Materials

Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

M-210-B4 cells (ATCC #CRL-1972) maintained in RPMI-1640 (Invitrogen, cat. no. 10-404-CV) supplemented with 10% fetal bovine serum (FBS; Hyclone, cat. no. SH30088.03) and 1% penicillin-streptomycin (Invitrogen, cat. no. 15140-122) Mitomycin C (American Pharmaceutical Partners, cat. no. 109020) divided into 900-μl aliquots and stored at −80◦ C M210-conditioned medium (see recipe) 0.1% (w/v) gelatin in distilled, deionized water and autoclaved for sterility Dulbecco’s phosphate-buffered saline (DPBS) without Ca+ and Mg2+ (CMF-DPBS) 0.05% (w/v) trypsin/0.53 mM EDTA Undifferentiated H1 and/or H9 human embryonic stem cells (WiCell; Kaufman et al., 2001) maintained on mouse embryonic fibroblast feeder cells Collagenase type IV (see recipe) ES cell wash medium (see recipe) for rinsing Differentiation medium (see recipe) for coculture 6-well tissue culture plates 15-ml conical tube 5-ml and 10-ml glass pipets (VWR, cat. nos. 53283-738 and 53283-740)

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Figure 1F.6.1 Overview of coculture strategy. Major steps in differentiation and CD34 selection using hES cell and stromal cell coculture system.

Inactivate M210 stromal cells and set up plates 1. To inactivate M210 stromal cells, incubate with 10 μg/ml mitomycin C in M210conditioned medium for 3 hr at 37◦ C with 5% CO2 . We typically use 21 to 23 ml of mitomycin C–treated M210-conditioned medium per 75-cm2 flask of M210 cells. Use of M210 “conditioned medium” by removing and saving medium from M210 cell culture is effective for this short incubation period and provides more economical use of reagents.

2. While mitomycin treatment of stromal cells is being done, coat the desired number of 6-well plates with 0.1% gelatin (2 ml/well) for at least 30 min prior to use. Aspirate gelatin immediately prior to adding inactivated stromal cells. 3. After incubation, remove the mitomycin C–containing medium, wash the cells twice, each time with CMF-DPBS (25 ml/flask per wash). CAUTION: Mitomycin C is a known teratogen. Handle with caution and dispose of according to standard institutional protocols. All DPBS used in wash steps must also be treated as hazardous waste and disposed of in the same manner. Alternatively, inactivation of stromal cells can be accomplished with irradiation treatment. Appropriate dose should be optimized based on which cell line is being utilized.

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4. Detach cells using 0.05% (w/v) trypsin/0.53 mM EDTA. When cells become nonadherent, wash the flask with 10 ml M210-conditioned medium, collect cells in a 15-ml conical tube, and centrifuge for 5 min at ∼425 × g, 4◦ C. Discard supernatant, resuspend pellet in 10 ml medium per 75-cm2 flask that was harvested, and count cells (UNIT 1C.3). 5. Set up confluent M210 monolayers on gelatin-coated 6-well plates by using 2.5 ml M210 medium containing 1.5 × 105 cells per ml for each well (2.25 × 106 cells in 15 ml total for each 6-well plate). Allow cells to attach overnight before continuing coculture procedure. When culturing cells in 6-well plates, care should be taken to ensure cells are evenly distributed throughout the well and do not settle in one area. Cells should be evenly distributed in the wells by gently shaking the plate (in two directions) on a flat surface after adding cells. This also prevents the cells from settling in clumps. Four nearly confluent 75-cm2 flasks of M210 cells can be used to generate ∼ten to fifteen 6-well plates.

Set up coculture 6. To disrupt undifferentiated human ES cell colonies, incubate cells with 1.5 ml per well of 1 mg/ml type IV collagenase for 5 to 10 min at 37◦ C with 5% CO2 . Observe cells under a microscope at 5-min intervals. Continue collagenase treatment until edges of colonies begin to lift off the plate. The passage of hES cells onto stromal cells is much like passage onto fresh MEFs for routine maintenance of undifferentiated hES cell culture. We would expect other hES cell lines to also give similar results in this system, though most of our experience is with H1 and H9 cells.

7. When colonies are starting to lift off, using a 10-ml glass pipet held at a right angle to the culture well with the tip flat, scrape the entire well and place cells in a 15-ml conical tube. Rinse the wells with additional ES cell wash medium as necessary to collect as many cells as possible. 8. Centrifuge cells for 4 min at 200 × g, 4◦ C. Gently wash the cell pellet once more using 6 to 9 ml ES cell wash medium. 9. Aspirate medium from M210 plates and add 1.5 ml differentiation medium to each well. Resuspend undifferentiated ES cells in 6 ml differentiation medium and add 1 ml cell solution to each well. Culture in 37◦ C with 5% CO2 for desired number of days with medium changes every 2 to 3 days, or more often if desired. One confluent 6-well plate of hES cell colonies is adequate to plate onto three to four 6-well plates of inactivated M210 stromal cells (passed at ∼1:3 ratio). Colony size and density determines the appropriate number of plates.

BASIC PROTOCOL 2

Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

PRODUCTION OF SINGLE-CELL SUSPENSION OF DIFFERENTIATED hES CELLS AND SELECTION OF HEMATOPOIETIC PROGENITORS VIA MAGNETIC SORTING It has been found that hES cells cocultured with M210 cells for a period of 7 to 21 days effectively produce reasonable numbers of hematopoietic progenitor cells. To determine the optimal number of days for coculture, a time course should be conducted and cells should be assayed every 2 to 3 days. However, after a prolonged period of coculture, the differentiated hES cells can become very dense. Therefore, production of a single-cell suspension from this heterogeneous cell population as needed for subsequent analyses is not a trivial exercise. The method described here is highly efficient with good cell viability and few cells lost during harvest and sorting.

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Figure 1F.6.2 Photographic representation of hES cells cocultured with M210 cells. Day 1 through 21 photographs representative of morphology changes and growth patterns of hES cells on M210 stromal cells during differentiation. Original magnification 20× (top rows) and 100× (bottom rows).

After the desired number of days in coculture, H9 cells can be harvested and sorted using the EasyStep magnetic system (or other magnetic system, as desired). Isolated CD34+ cells can be analyzed via flow cytometry and placed in hematopoietic culture conditions for further differentiation and growth. In our experience, CD34+ cells begin to appear after ∼3 to 5 days of stromal cell coculture, and CD45+ cells appear at roughly day 14. Typically, the maximum number of CD34+ CD45+ double positive cells emerge during later time points (day 18 to 21; Kaufman et al., 2001; Tian et al., 2006). Development of colony-forming cells (CFCs) closely parallels development of CD34+ CD45+ cells, as previously described (Kaufman et al., 2001; Woll et al., 2008). Gene expression analyses can also be done by RT-PCR from either unsorted cell populations, or specific populations of phenotypically sorted cells during this time course, as previously demonstrated (Kaufman et al., 2001; Tian et al., 2006; Woll et al., 2008). Figure 1F.6.2 provides representative photos of hES cell differentiation via stromal cell coculture.

Materials hESCs cocultured with preferred stromal cell line (e.g., M210; see Basic Protocol 1) Collagenase type IV (see recipe) ES cell wash medium (see recipe) for rinsing

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Dulbecco’s phosphate-buffered saline (DPBS) without Ca+ and Mg2+ (CMF-DPBS) 0.05% (w/v) trypsin/0.53mM EDTA (Cellgro/Mediatech, cat. no. 25-052-CI) Chick serum (Sigma) EasySep buffer consisting of CMF-DPBS supplemented with 2% (v/v) FBS and 1 mM EDTA (stored at 2◦ to 8◦ C) EasySep human CD34 selection kit (StemCell Technologies, cat. no. 18056) containing: CD34 positive selection cocktail Magnetic nanoparticle solution FACS buffer (see recipe) Hematopoietic culture medium of choice Appropriate antibodies Appropriately labeled isotype controls Propidium iodide or 7-AAD 10-ml glass pipets 15-ml conical tubes 37◦ C water bath Vortex 100-μm cell filter/strainer (Partec CellTrics, cat. no. 04-0042-2318) FACS tube, sterile EasySep Magnet (StemCell Technologies, cat. no. 18000) Additional reagents and equipment for counting cells (UNIT 1C.3) Generate a single-cell suspension 1. Remove the culture medium and disrupt cells using 1.5 ml collagenase type IV solution (1 mg/ml) per well for 7 to 8 min at 37◦ C with 5% CO2 . Then add 1.5 ml ES cell wash medium. 2. Using a 10-ml glass pipet, scrape the cells (as described in Basic Protocol 1, step 7) and collect the cells in conical tubes (2 to 3 wells/15-ml conical tube). Wash each well with 1.5 to 2.0 ml ES cell wash medium and add to the same conical tube as the cells. 3. Centrifuge cells for 4 min at 425 × g, 4◦ C and wash the cell pellet with 10 ml CMF-DPBS; centrifuge again using the same conditions. 4. Remove the supernatant and dissociate the cell clumps by incubating with 2 ml/tube 0.05% trypsin/EDTA containing 2% chick serum in a water bath 5 to 15 min at 37◦ C. Manually shake and vortex the tubes at regular intervals (approximately every 5 min) throughout the incubation to improve the quality of the single-cell suspension. Incubation can be continued as needed until few cell clumps can be visualized, though >15 min is not recommended, as it is unlikely that better separation of cells will be seen after this time, and viability may be decreased. Adding 2% chick serum to the trypsin digestion provides protein bulk during this incubation step, but unlike FBS, chick serum does not contain trypsin inhibitors.

5. Add 10 ml ES cell wash medium to the digestion tube and pipet the solution up and down. Centrifuge 4 min at 425 × g, 4◦ C. Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

6. Aspirate the supernatant and resuspend cells in ∼3 ml ES cell wash medium. Filter cells into a new 15- or 50-ml conical tube using the 100-μm cell strainer. Set aside a small aliquot of this cell population for presort flow cytometric analysis and count cells (UNIT 1C.3). A 50-μm CellTrics strainer can also be utilized in this step.

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Isolate CD34+ cells 7. Move all cells to a sterile FACS tube and centrifuge 5 min at 425 × g, 4◦ C. Wash cells in ∼2.5 ml of EasySep buffer (sterile) and centrifuge again. Keep EasySep buffer on ice (or at 4◦ C) throughout procedure.

8. Resuspend cells in EasySep buffer as indicated in the EasySep protocol (for 2 × 107 cells, use 1 ml, for <2 × 107 cells, use 100 to 500 μl). 9. Add the EasySep CD34+ selection cocktail to the cell suspension for 15 min at the concentration indicated (1 μl cocktail per 10 μl of cell suspension) and pipet up and down to mix. Other magnetic separation systems appropriate for this procedure are commercially available. The EasySep system has been used almost exclusively in our practice.

10. Next, add the EasySep magnetic nanoparticle solution to the cell suspension for 10 min. Add half of the amount of cocktail used and pipet up and down to mix. 11. Before placing the tube into the magnet, add ∼2.5 ml of EasySep buffer and pipet up and down to mix. Place the FACS tube into the magnet for 6 min and do not disrupt surrounding area during this time. Carefully decant the EasySep buffer without moving the FACS tube. Often there is a residual drop at the opening of the tube after decanting, do not attempt to shake or blot this drop from the tube, allow it to remain.

12. Repeat this wash step two more times. 13. Resuspend cells in hematopoietic culture medium of choice and remove a small aliquot (∼100 μl) of cells to count (UNIT 1C.3). Remove another aliquot of cells for post-sort flow cytometric analysis to assess purity of CD34+ cells and other hematopoietic cell characteristics.

Perform flow cytometric analysis 14. To wash the presort and post-sort cell populations, add 2.5 ml FACS buffer to each tube. Based upon the number of desired stains, divide the cells into an appropriate number of tubes (2.0 × 104 to 3.0 × 104 cells per tube for FACS) and centrifuge 5 min at 425 × g in a temperature-controlled unit (keep temperature at 4◦ C). 15. Decant and add antibodies to analyze cellular characteristics. Also, include appropriately labeled isotype controls to establish machine settings and background signal

Figure 1F.6.3 Flow-cytometric analysis of hES cells differentiated by coculture with M210 cells. (A) Cell population analyzed for CD34 and CD45 prior to magnetic sorting for CD34. These hES cells were cocultured on M210 stromal cells for 19 days. (B) The identical cell population sorted for CD34+ cells and re-analyzed to demonstrate enrichment of CD34+ CD45+ cells.

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levels. Incubate 30 min at 4◦ C and wash twice, each time with 2 ml FACS buffer and centrifuge 5 min at 425 × g, 4◦ C. Suggested hematopoietic antibody markers include CD34, CD45, c-kit, BB9, and CD56.

16. Decant and add ∼200 μl of FACS buffer to each tube. Label to exclude dead cells by adding 1.5 μl propidium iodide or 7-AAD and vortex briefly. Perform analysis using standard methods. Figure 1F.6.3 illustrates typical flow cytometric results generated from these methods. SUPPORT PROTOCOL 1

CULTURE OF hES CELLS hES cells must be maintained in culture for undifferentiated cell growth before culturing them with stromal cells for inducing differentiation.

Additional Materials (see Basic Protocol 1) hES cells hES cell medium (see recipe) for undifferentiated cell growth Undifferentiated hES cells are maintained in culture as previously described (Kaufman et al., 2001). Briefly, undifferentiated hES cells are passed to new plates of MEFs approximately every 5 to 7 days using methods similar to passing undifferentiated hESCs onto stromal cells to initiate the differentiation culture, as described in steps 5 to 7 of Basic Protocol 1 in the coculture set up instructions above. Grown on feeder plates consisting of either mouse embryonic fibroblasts (MEFs) or Matrigel-coated plates, cells are maintained in colony form and given fresh medium daily. Cells are typically passed to new plates every 4 to 6 days to prevent differentiation and maintain pluripotency. SUPPORT PROTOCOL 2

CULTURE OF M210 CELLS M210 cells are maintained for use as stromal cells in cocultures with hES cells to induce differentiation. For optimal differentiation, M210 stromal cells should be discarded after being passaged for roughly 3 months (∼25 passages) and a fresh stock of earlier passage cells should be used. As noted above, we have found that M210 cells are more stable in their hematopoietic supporting potential than S17 cells, which require use of fresh cells approximately every 1 to 2 months.

Materials 0.1% (w/v) gelatin in distilled deionized water and autoclaved for sterility M210 stromal cells M210 culture medium (see recipe) Dulbecco’s phosphate-buffered saline (DPBS) without Ca+ and Mg2+ (CMF-DPBS) 0.05% (w/v) trypsin/0.53 mM EDTA 75-cm2 flasks 10-ml glass pipets 15-ml conical tubes 1. Prepare 75-cm2 flask (three for each flask to be split) by adding 7 ml of 0.1% gelatin at least 30 min prior to use.

Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

2. Aspirate M210 culture medium from a M210 culture in a 75-cm2 flask and rinse flask with 10 ml CMF-DPBS. Add 2 ml trypsin/EDTA and incubate at 37◦ C with 5% CO2 . 3. When cells become nonadherent, add 10 ml M210 culture medium and using a 10-ml glass pipet, scrape the cells (as described in Basic Protocol 1, step 7) and collect cells

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into a 15-ml conical tubes (one 75-cm2 flask per tube). Centrifuge 4 min at 425 × g, 4◦ C. 4. During centrifugation, begin preparing new 75-cm2 flasks by aspirating the 0.1% gelatin and adding 14 ml M210 medium to each flask. 5. Aspirate the supernatant from the 15-ml conical tubes and resuspend the cells in 6 ml M210 medium. Add 2 ml of the cell suspension to each of the new flasks. Gently shake the culture flask on a flat surface and return it to the incubator. 6. Passage the cultures every 3 to 4 days, before they reach confluence.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Collagenase type IV Dilute collagenase type IV (Invitrogen, cat. no. 17104-019) to a concentration of 1 mg/ml in DMEM/F12. Filter sterilize with a 0.22-μm Steriflip membrane (Millipore, cat. no. SCGP00525) into a 50-ml conical tube. Store up to 7 days at 4◦ C.

Differentiation medium RPMI-1640 supplemented with: 15% (v/v) defined fetal bovine serum (FBS; Hyclone, cat. no. SH30070.03) 1% (v/v) penicillin/streptomycin 2 mM L-glutamine (see recipe) 0.2% (v/v) 2-mercaptoethanol 1% (v/v) non-essential amino acids Filter sterilize Store up to 7 to 10 days at 4◦ C Defined FBS lots are individually tested to determine ability to support induction of differentiation.

ES cell wash medium DMEM/F12 (Invitrogen, cat. no. 11330-032) supplemented with: 10% (v/v) knock-out serum replacement (Invitrogen, cat. no. 10828-028) 1% (v/v) penicillin/streptomycin Filter sterilize Store up to 6 months at 4◦ C FACS buffer 2% (v/v) standard FBS Dulbecco’s phosphate-buffered saline (DPBS) without Ca+ and Mg2+ (CMFDPBS) Store up to 6 months at 4◦ C This solution does not need to be filter sterilized.

hES cell medium DMEM/F12 (Invitrogen, cat. no. 11330-032) supplemented with: 15% (v/v) knock-out serum replacement (Invitrogen, cat. no. 10828-028) 1% (v/v) penicillin/streptomycin 1% (v/v) non-essential amino acid solution (Invitrogen, cat. no. 11140-050) 4 ng/ml basic fibroblast growth factor (Invitrogen, cat. no. 13256)

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1% (v/v) L-glutamine (see recipe; 2 mM final) Filter sterilize Store up to 7 days at 4◦ C L-glutamine

Mix 0.146g L-glutamine powder with 10 ml CMF-DPBS and 7 μl 2-mercaptoethanol. Store up to 7 days at 4◦ C.

M210 medium RPMI-1640 supplemented with: 10% (v/v) standard fetal bovine serum (for cell culture) 1% (v/v) penicillin/streptomycin Store up to 7 days at 4◦ C COMMENTARY Background Information

Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

One of the benefits of studying hES cells is the ability to dissect out distinct mechanisms that regulate human development and analyze each process in depth on a cell-specific basis. An accurate in vitro model system can illuminate not only normal differentiation pathways but also where potential errors occur. With this new insight, enhanced differentiation models can be potentially utilized to generate multipotent hematopoietic progenitors quickly and more efficiently. Such knowledge will be useful for time-sensitive therapeutic applications, as well as generating an accessible stock source of cells for cytotoxicity and drug studies. As discussed, differentiation of hES cells can be accomplished via stromal cell coculture, as well as through embryoid body formation. Because of the ability to create specific cell culture conditions in a more controlled manner via coculture, it is possible to generate distinct hematopoietic cell lineages in a step-wise fashion. The initial coculture conditions with defined serum is the first step. Further culture of sorted CD34+ cells in lineagespecific conditions, using cocktails of defined cytokines, typically yields more homogeneous cell populations of the lineage of choice. For example, specific myeloid, erythroid, and lymphoid cells have been derived and maintained in culture (Woll et al., 2005; Anderson et al., 2006; Olivier et al., 2006). One goal of hES cell-based hematopoietic differentiation is to create cells capable of repopulating an individual who has a given blood disorder. Initial attempts with transplantation of hES cell-derived blood cells transplanted into immunodeficient mice or fetal sheep model have had only modest success (Zanjani, 2000; Wang et al., 2005; Narayan

et al., 2006; Tian et al., 2006). While hES cellderived cells are able to survive after transplantation and demonstrate long-term engraftment, the level of engraftment has been modest, and the range of differentiated cells that develop in vivo is typically limited. Notably, a recent study compared differentiation of hES cells on different mouse stromal cell lines and found that coculture with AM20.1B4 cells lead to improved hematopoietic engraftment (Ledran et al., 2008). However, in our studies, the differentiated hES cells did not exhibit teratoma formation (a common concern), even when transplanted into immunodeficient mice given additional radiation and immune-depletion via anti-NK cell antibodies (Tian et al., 2006).

Critical Parameters and Troubleshooting The quality of hematopoietic differentiation is dependant upon a number of factors. The initial state of the hES cells used in the study is of utmost importance. Maintenance of the cells in an undifferentiated state is a critical and labor-intensive process. Mastering the requirements to care for hES cells takes time, effort, and experience. Observation and feeding of hES cells on a daily basis must be made in order to monitor their growth rate and degree of differentiation. By watching the size and shape of colonies over time, one can accurately gauge the quality of the cells and determine when they require passaging. Another important factor in hES cell culture is the state of the MEF feeder cells. Initial inactivation and proper plating density determines how well the hES cells will replicate in an undifferentiated state. If MEF cells are plated at incorrect densities, spontaneous differentiation is more likely to occur. MEFs should be plated at a density at which cells just contact

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one another, but do not overlap. This density varies by MEF source (usually between 1.875 to 2.25 × 105 cells per well) hES cells which have started differentiating cannot be used in further hematopoietic differentiation studies, as a poor yield of cells with appropriate characteristics will be obtained. The process of setting up differentiation studies also takes time and experience to learn. Without obtaining exact hES cell counts prior to setting up an experiment (cells cannot be in a single-cell suspension prior to plating for differentiation), a researcher must develop an idea of appropriate plating density rather than abiding by established protocol numbers. If the stromal cell layer is plated too dense, the hES cells will not have space to grow and differentiate, causing a low yield of CD34+ cells. Initial plating density of undifferentiated hESCs (onto M210s) should mimic the density at which the colonies are passaged onto feeder cells. Complete inactivation of stromal cells must be achieved, otherwise subsequent cell growth will inhibit proper differentiation. If poor differentiation results are noted with complete inactivation of stromal cells and proper plating densities, it may be best to test another lot of defined serum. Medium used for differentiation culture should be prepared fresh every 7 to 10 days. Although medium is stored at 4◦ C, it should be allowed to warm to room temperature before use during culture procedures. While harvesting the cells at the end of the differentiation period, there are several steps that should be executed with care. When generating the single-cell suspension, it is helpful to add the 2% chick serum. This improves cell viability and does not contain the trypsin inhibitors present in many other types of serum. After the suspension is generated, it is important to wash the cell filters to obtain maximum yield. Presort cell populations <8 × 106 are difficult to work with and often do not give great numbers of CD34+ cells. During magnetic sorting, some cells are inevitably lost, even if they are CD34+ . To minimize loss, the magnetic-sorting apparatus should not experience vibrations or other disruptions while the tube and labeled cells are in place. When decanting the tube during wash steps, it is also worth noting that the liquid remaining at the end of the tube should not be blotted or shaken from it, but rather left in place. As always, sterile technique should be maintained throughout the procedure. Research of known developmental pathways in hematopoiesis should also be con-

ducted in order to select proper cytokines and growth conditions for CD34+ cells. Again, this will aid in obtaining specific mature cell lineages.

Anticipated Results After hES cells are plated onto inactivated M210 stromal cell layers, they will appear as round clumps, similar to when they are first passed onto MEF plates for regular maintenance. In the early days after initial plating, it is difficult to view the progress of smaller colonies. As the days progress, clumps will grow and spread out onto the plate as they begin to display characteristic irregular edges of differentiating colonies. During later days, colonies will begin to contact one another and blend together. The efficiency of differentiation will depend upon initial hES cell quality, the type of stromal cell layer implemented in coculture, cytokine and growth media selection, and number of days in culture. When setting up coculture experiments with new stromal cell lines, differentiation characteristics of the cells should be monitored on a time course via flow cytometry. As expression levels of various markers rise and fall over time, it is possible to map the process by which hematopoietic precursors develop. Comparisons can also be made between presorted and post-sorted cells. By assessing these two differentiated populations, further conclusions can be drawn about which markers are co-expressed and to what degree. After this data is collected, it is possible to optimize results based on desired progenitor type. Initial unsorted differentiated cell populations can exhibit upwards of 15% CD34+ cells (Tian et al., 2006). After sorting for CD34, populations can reach up to 95% purity. Coexpression of CD34 and other hematopoietic markers fluctuates by number of days in coculture and quality of differentiation. After 20 days of coculture-mediated differentiation, CD45 expression is typically 1% to 5% of the total differentiated cell population, and >20% of the sorted CD34+ cell population.

Time Considerations It is best to coordinate undifferentiated hES cell culture and setting up differentiation studies. hES cells should be passaged onto new feeder cells every 5 to 7 days, depending on the quality of MEFs (ability to sustain undifferentiated cells). New MEF plates should be prepared at least 1 day prior to their use but

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can be used for up to 6 to 7 days. Appropriate MEF density, which is subconfluent, varies from batch to batch. Setting up inactivated stromal cell plates the day prior to a day hES cells require passage is optimal, although inactivated stromal cell plates can be used for 2 to 3 days after plating. Stromal cell culture requirements vary from line to line. As a general rule, these cells should be passaged every 3 to 4 days. Depending on the number of plates, setting up hES cells on stromal cell plates will require ∼1 to 2 hr. Harvesting, sorting, and analyzing the cells after differentiation is more time-intensive, requiring 2 to 4 hr to complete.

Literature Cited Anderson, J.S., Bandi, S., Kaufman, D.S., and Akkina, R. 2006. Derivation of normal macrophages from human embryonic stem (hES) cells for applications in HIV gene therapy. Retrovirology 3:24. Chadwick, K., Wang, L., Li, L., Menendez, P., Murdoch, B., Rouleau, A., and Bhatia, M. 2003. Cytokines and BMP-4 promote hematopoietic differentiation of human embryonic stem cells. Blood 102:906-915. Kaufman, D.S., Hanson, E.T., Lewis, R.L., Auerbach, R., and Thomson, J.A. 2001. Hematopoietic colony-forming cells derived from human embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A 98:10716-10721. Keller, G. 2005. Embryonic stem cell differentiation: Emergence of a new era in biology and medicine. Genes Dev. 19:1129-1155. Kennedy, M., D’Souza, S.L., Lynch-Kattman, M., Schwantz, S., and Keller, G. 2007. Development of the hemangioblast defines the onset of hematopoiesis in human ES cell differentiation cultures. Blood 109:2679-2687. Ledran, M.H., Krassowska, A., Armstrong, L., Dimmick, I., Renstrˆm, J., Lang, R., Yung, S., Santibanez-Coref, M., Dzierzak, E., Stojkovic, M., Oostendorp, R.A., Forrester, I., and Lake, M. 2008. Efficient hematopoietic differentiation of human embryonic stem cells on stromal cells derived from hematopoietic niches. Cell Stem Cell 3:85-99. Mummery, C., Ward-van Oostwaard, D., Doevendans, P., Spijker, R., van den Brink, S., Hassink, R., van der Heyden, M., Opthof, T., Pera, M., de la Riviere, A. B., Passier, R., and Tertoolen, I. 2003. Differentiation of human embryonic stem cells to cardiomyocytes: Role of coculture with visceral endoderm-like cells. Circulation 107:2733-2740.

Hematopoietic Differentiation of hESC by Cocultivation with Stromal Layers

Mummery, C., van der Heyden, M.A., deBoer, T.P., Passier, R., Ward, D., van den Brink, S., van Rooijen, M., and van de Stolpe, A. 2007. Cardiomyocytes from human and mouse embryonic stem cells. Methods Mol. Med. 140:249272.

Narayan, A.D., Chase, J.L., Lewis, R.L., Tian, X., Kaufman, D.S., Thomson, J.A., and Zanjani, E.D. 2006. Human embryonic stem cell-derived hematopoietic cells are capable of engrafting primary as well as secondary fetal sheep recipients. Blood 107:2180-2183. Olivier, E.N., Qiu, C., Velho, M., Hirsch, R.E., and Bouhassira, E.E. 2006. Large-scale production of embryonic red blood cells from human embryonic stem cells. Exp. Hematol. 34:16351642. Schmitt, T.M., de Pooter, R.F., Gronski, M.A., Cho, S.K., Ohashi, P.S., and Zuniga-Pflucker, J.C. 2004. Induction of T cell development and establishment of T cell competence from embryonic stem cells differentiated in vitro. Nat. Immunol. 5:410-417. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Tian, X. and Kaufman, D.S. 2008. Hematopoietic development of human embryonic stem cells in culture. In Hematopoietic Stem Cell Protocols (K.D. Bunting, ed.) pp. 119-133. Humana Press, Totowa, N.J. Tian, X., Woll, P.S., Morris, J.K., Linehan, J.L., and Kaufman, D.S. 2006. Hematopoietic engraftment of human embryonic stem cell-derived cells is regulated by recipient innate immunity. Stem Cells 24:1370-1380. Vodyanik, M.A., Bork, J.A., Thomson, J.A., and Slukvin, I.I. 2005. Human embryonic stem cellderived CD34+ cells: Efficient production in the coculture with OP9 stromal cells and analysis of lymphohematopoietic potential. Blood 105:617626. Wang, L., Menendez, P., Shojaei, F., Li, L., Mazurier, F., Dick, J.E., Cerdan, C., Levac, K., and Bhatia, M. 2005. Generation of hematopoietic repopulating cells from human embryonic stem cells independent of ectopic HOXB4 expression. J. Exp. Med. 201:1603-1614. Woll, P.S., Martin, C.H., Miller, J.S., and Kaufman, D.S. 2005. Human embryonic stem cell-derived NK cells acquire functional receptors and cytolytic activity. J. Immunol. 175:5095-5103. Woll, P.S., Morris, J.K., Painschab, M.S., Marcus, R.K., Kohn, A.D., Biechele, T.L., Moon, R.T., and Kaufman, D.S. 2008. Wnt signaling promotes hematoendothelial cell development from human embryonic stem cells. Blood 111:122131. Zanjani, E.D. 2000. The human sheep xenograft model for the study of the in vivo potential of human HSC and in utero gene transfer. Stem Cells 18:151. Zeng, X., Cai, J., Chen, J., Luo, Y., You, Z.-B., Fotter, E., Wang, Y., Harvey, B., Miura, T., Backman, C., Chen, G.J., Rao, M.S., and Freed, W.J. 2004. Dopaminergic differentiation of human embryonic stem cells. Stem Cells 22:925940.

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TLX1 (HOX11) Immortalization of Embryonic Stem Cell–Derived and Primary Murine Hematopoietic Progenitors

UNIT 1F.7

Robert G. Hawley,1 Teresa S. Hawley,2 and Alan B. Cantor3, 4 1

The George Washington University Medical Center, Washington, D.C. Flow Cytometry Core Facility, The George Washington University Medical Center, Washington, D.C. 3 Department of Pediatric Hematology-Oncology, Children’s Hospital Boston, Boston, Massachusetts 4 Dana-Farber Cancer Institute, Harvard Medical School, Boston, Massachusetts 2

ABSTRACT The ability to generate genetically engineered cell lines is of great experimental value. They provide a renewable source of material that may be suitable for biochemical analyses, chromatin immunoprecipitation assays, structure-function studies, gene function assignment, and transcription factor target gene identification. This unit describes protocols for TLX1 (HOX11)-mediated immortalization of murine hematopoietic progenitors derived from in vitro differentiated murine embryonic stem cells, or from primary mouse fetal liver or bone marrow. A wide variety of hematopoietic cell types have been immortalized using these procedures including erythroid, megakaryocytic, monocytic, myelocytic, and multipotential cell types. These lines are typically cytokine dependent C 2008 by for their survival and growth. Curr. Protoc. Stem Cell Biol. 7:1F.7.1-1F.7.19.  John Wiley & Sons, Inc. Keywords: murine cell immortalization r TLX1 r HOX11 r hematopoietic in vitro ES cell differentiation r hematopoietic progenitors

INTRODUCTION The ability to disrupt and/or manipulate selected genes in embryonic stem (ES) cells and whole animals has revolutionized the study of molecular and developmental biology. However, primary tissue from gene-targeted mice has limited applications because of the relatively small amount of material available, heterogeneity of cell types present, variability due to harvests from multiple animals, and cumbersome nature of accessing material for repeated experiments. Generation of immortalized cell lines from in vitro differentiated gene-targeted ES cells or primary cells from gene-targeted animals allows for a uniform, self-sustaining source of material for repeated studies. Several different approaches have been reported for the immortalization of primary murine hematopoietic cells that include retroviral expression of Myc (Green et al., 1989), Myb (Gonda et al., 1989), Hoxb8 (Hox-2.4; Perkins and Cory, 1993), TLX1 (formerly called HOX11; Hawley et al., 1994a), E2A-Pbx1 (Kamps and Wright, 1994), MLL (Lavau et al., 1997), Lhx2 (Pinto do et al., 2002), RARA (Du et al., 1999), Hoxa9 (Calvo et al., 2000; Schnabel et al., 2000), Notch1 (Varnum-Finney et al., 2000), v-raf/v-myc (Coghill et al., 2001), MYST3-NCOA2 (Deguchi et al., 2003), Evi1 (Du et al., 2005), HOXB6 (Fischbach et al., 2005), HOXB4 (Zhang et al., 2007), β-catenin (Templin et al., 2008), and Id1 (Suh et al., 2008). A subset of the genes utilized in these studies—most notably, TLX1—have also been demonstrated to efficiently immortalize ES cell–derived Current Protocols in Stem Cell Biology 1F.7.1-1F.7.19 Published online December 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f07s7 C 2008 John Wiley & Sons, Inc. Copyright 

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hematopoietic cells (Keller et al., 1998; Cantor et al., 2002; Shaw et al., 2006; Riz et al., 2007). The protocols in this unit can be used to provide progenitor cells that are rare, transient, and otherwise difficult to purify, for further characterization and study. The following protocols were developed for retroviral TLX1 immortalization of hematopoietic progenitors derived from in vitro differentiated murine ES cells (Basic Protocol) as well as from primary murine hematopoietic tissues (Alternate Protocol). Support Protocol 1 addresses the maintenance of TLX1 retroviral producer cell lines. Support Protocols 2, 3, and 4 (respectively) address cloning, freezing, and characterizing of the immortalized cells. CAUTION: TLX1 is a well characterized human oncogene. The protocols described in this unit involve retroviral transduction of TLX1 into murine cells. The retroviral particles are designed to be replication-defective and are generated in murine ecotropic retroviral producer cell lines. Nonetheless, caution should be taken when handling the retroviral supernatants. Standard microbiologic safety procedures (see http://www.absa.org/restraining.html) should be followed including the use of gloves, lab coats, and safety goggles when handling the retroviral supernatants. All materials and surfaces coming in contact with the retroviral supernatants should be decontaminated following standard procedures. The protocols described below should not be altered. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used throughout. NOTE: All incubations are performed in a humidified 37◦ C/5% CO2 tissue culture incubator unless stated otherwise. BASIC PROTOCOL

TLX1 IMMORTALIZATION OF ES CELL IN VITRO DIFFERENTIATED HEMATOPOIETIC PROGENITORS This protocol involves in vitro differentiation of murine ES cells to embryoid bodies (EBs) under conditions that promote hematopoietic development. The EBs (containing hematopoietic progenitor cells) are then disaggregated and the released cells transduced with TLX1-expressing retroviruses by coculture with TLX1 retroviral producer cells to generate immortalized hematopoietic cells. Single cell clones are then isolated to produce cell lines.

Materials

TLX1 Immortalization of Hematopoietic Progenitors

Murine ES cells, gel-adapted (grown on gelatinized plates, not on feeder cells) and of low passage number (see UNIT 1C.4) IMDM-ES-15 with MTG, LIF, and pen/strep (see recipe) 0.05% (w/v) and 0.25% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25200 and 25300, respectively) IMDM-ES-5 with MTG and pen/strep (see recipe) Primary ES cell differentiation medium (see recipe) TLX1-retroviral producer cell line (see Support Protocol): GP+E-86/MSCV-HOX11 for cotransduction of the neomycin phosphotransferase (neo) gene conferring resistance to the neomycin analog Geneticin in mammalian cells or GP+E-86/MSCVhyg-HOX11 for cotransduction of the hygromycin B phosphotransferase (hyg) drug resistance gene Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) IMDM-ES-15: IMDM supplemented with 15% (v/v) ES-FBS

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Coculture medium (see recipe) IMDM-ES-15 with glutamine and pen/strep (see recipe) Immortalized cell medium (see recipe) 50 mg/ml Geneticin (Invitrogen cat. no. 10131-035) or 50 mg/ml hygromycin B (Mediatech) stock Gelatinized 25-cm2 tissue culture flasks (see recipe) 15-ml and 50-ml conical centrifuge tubes (sterile) Tissue culture centrifuge (refrigerated, benchtop centrifuge with swinging bucket rotor) 100-mm petri dishes 100-mm and 60-mm tissue culture dishes 20-G needle and 3-ml syringe Hemacytometer Cell irradiator (optional) Additional reagents and equipment for counting viable cells (UNIT 1C.3) Prepare ES cells 1. Two days before inducing differentiation, split the murine gel-adapted ES cells at various dilutions (e.g., 1:10, 1:5, 1:3) into 10 ml of IMDM-ES-15 with MTG, LIF, and pen/strep and plate in gelatinized 25-cm2 flasks. Refeed cells daily. 2. After two days, choose a flask with cells at ∼25% to 50% confluency. Wash cells briefly with 2 ml of 0.05% trypsin/EDTA, and then add 2 ml of 0.25% trypsin/EDTA. Incubate until cells start to loosen from the flask (about 3 min). 3. Pipet up and down several times to dissociate cells and add the suspension to 10 ml of IMDM-ES-15 with MTG, LIF, and pen/strep in a 15-ml centrifuge tube. 4. Centrifuge 5 min at ∼200 × g, 4◦ C in a tissue culture centrifuge. 5. Carefully remove the supernatant and resuspend the pellet in 5 ml of IMDM-ES-5 with MTG and pen/strep. 6. Count the number of viable cells in a hemacytometer (UNIT 1C.3).

Differentiate ES cells to EBs 7. Add 50,000 cells to each of three 100-mm petri dishes (do not use tissue culture dishes) containing 10 ml primary ES differentiation medium. The cells will probably need to be diluted 1:10, using IMDM-ES-5 with MTG and pen/strep. IMDM is used for dilutions here and in step 5 (instead of differentiation medium) in order to minimize waste of cytokines, which are expensive. It is important to use petri plates (which are not coated) in this step because the ES cells will not form EBs on tissue culture dishes (which are coated with a material, usually poly-lysine, to promote cell attachment).

8. Incubate for 6 or 7 days without changing the medium. The length of incubation will depend on stage of differentiation and cell type targeted for immortalization. We have noted lower immortalization efficiency from EBs differentiated for fewer than 6 days.

9. During the EB differentiation, grow TLX1 retroviral producer cells in 100-mm tissue culture dishes, aiming to have cells at ∼90% to 100% confluency on the day of coculture with the ES-derived hematopoietic progenitors (day 6 to 7 of EB differentiation). Typically, a confluent dish of producer cells split ∼1:7 to 1:10 will be 90% to 100% confluent in ∼3 days.

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Dissociate EBs 10. At the end of the EB differentiation period, combine the EBs and medium from the three dishes in a 50-ml conical tube. Centrifuge 5 min at ∼200 × g, 4◦ C. 11. Carefully aspirate and discard supernatant. Resuspend the EBs in 5 ml of IMDM and then allow them to settle by gravity for ∼5 min. 12. Aspirate supernatant and add 3 ml of 0.25% trypsin/EDTA to the pellet. Incubate 3 min at room temperature. 13. Dissociate embryoid body cells by gentle passage through a 20-G needle six times. 14. Immediately add the suspension to 10 ml of IMDM-ES-15 to neutralize trypsin. 15. Measure the concentration of viable cells using a hemacytometer (UNIT 1C.3). 16. Centrifuge 3 × 106 cells 5 min at ∼200 × g, 4◦ C, and resuspend in 10 ml coculture medium.

Prepare producer cells 17. Irradiate the TLX1 retroviral producer monolayer with 3 Gy. The retroviral producer cell monolayer can be irradiated earlier in the day before dissociation of the EBs, and the irradiated cells left in the incubator until the time of the coculture. Although irradiation of the producer cell line is recommended, it is not absolutely necessary because the immortalized hematopoietic cells grow in suspension and the producer cells are adherent. Therefore, any contaminating retroviral producer cells that get carried over after the coculture period can eventually be removed by serial passage of the populations on tissue culture plates. If gene-targeted cells containing a neo targeting construct are immortalized with the MSCV-HOX11 retrovirus, then any remaining producer cells can also be eliminated by selection in Geneticin. If the producer cell line is not irradiated, the plating density should be slightly lower (e.g., ∼50% to 70% confluent in a 100-mm tissue culture dish); otherwise, producer cells may become confluent and lift off the dish.

18. Wash the irradiated cell monolayer once with 5 ml of IMDM-ES-15 with glutamine and pen/strep.

Immortalize and passage EB cells 19. Gently add the disaggregated EB cell suspension to the irradiated cell monolayer. Incubate the coculture for 3 days without changing the medium. 20. After 3 days, gently agitate the dishes to remove suspension cells loosely adherent to the monolayer and transfer the supernatant to a 15-ml conical tube. 21. Centrifuge 5 min at ∼200 × g, 4◦ C. 22. Resuspend the cell pellet in 3 ml immortalized cell medium and incubate in 60-mm tissue culture dish. Over the next few days, clusters of proliferating cells in suspension should be visible. In our experience, TLX1 immortalized cells are almost always IL-3 dependent for their growth and survival. Therefore, it is critical to include IL-3 in the medium after the coculture and in all subsequent steps. However, the IL-3-containing medium can be supplemented with other cytokines/growth factors, e.g., SCF, Epo (Keller et al., 1998; Yu et al., 2002; Riz et al., 2007). TLX1 Immortalization of Hematopoietic Progenitors

23. Select for TLX1-expressing cells (step 24) as early as 48 to 72 hr following the coculture, or passage cells in immortalized cell medium with the selective agent at 1:10 to 1:20 every 3 to 4 days, with gentle pipetting up and down to disrupt cell clusters.

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Select for TLX1-expressing immortalized cells 24a. For cells transduced with the neo-carrying retrovirus cell line (GP+E-86/MSCVHOX11): Add Geneticin to immortalized cell medium to a final concentration of 400 μg/ml. 24b. For cells transduced with the hyg-carrying retrovirus cell line (GP+E86/MSCVhyg-HOX11): Add hygromycin B to immortalized cell medium to a final concentration of 400 μg/ml. While selection for retrovirally transduced cells with Geneticin or hygromycin B is recommended, it is not strictly required since TLX1-expressing cells will exhibit a growth advantage over nontransduced cells during in vitro passaging (Yu et al., 2002; Su et al., 2006) and nontransduced cells should eventually disappear from the cultures due to senescence. However, this may take as long as several months, especially for long-lived primary cells, e.g., mast cells.

25. Clone Geneticin- or hygromycin B-resistant cells in methylcellulose (as described in Support Protocol 2) or by limiting dilution in liquid culture (see Fuller et al., 2001) using immortalized cell medium containing the appropriate selective antibiotic. See Figure 1F.7.1 for an example of a TLX1 immortalized cell line derived from in vitro differentiated ES cells. The IL-3-containing methylcellulose and medium can be supplemented with other cytokines/growth factors, e.g., SCF, Epo (Keller et al., 1998; Yu et al., 2002; Riz et al., 2007).

Figure 1F.7.1 Photomicrographs of a TLX1 immortalized clonal hematopoietic cell line derived from in vitro differentiated FOG-1–/– ES cells. (A) Phase contrast photomicrograph of the suspension cell culture (original magnification 100×). (B) Bright field photomicrograph of May-GrunwaldGiemsa stained cytospun cells (original magnification 1000×). Note that the absence of FOG-1 contributes to the immature and uniform appearance of the cells.

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26. Freeze clonal cell lines as described in Support Protocol 3. 27. Characterize the clonal lines, depending on the particular application, e.g., by identifying the hematopoietic cell lineages represented, and their relative state of maturation (Support Protocol 4). ALTERNATE PROTOCOL

TLX1 IMMORTALIZATION OF PRIMARY MURINE HEMATOPOIETIC CELLS In this protocol, primary hematopoietic cells are harvested from murine fetal liver (FL) or bone marrow (BM) and then cocultured with TLX1-expressing retroviral producer cell lines. After coculture, cells are selected for retroviral expression and cloned to generate immortalized cell lines. Procedures for immortalization of yolk sac–derived precursors cells have also been published (Yu et al., 2002).

Additional Materials (also see Basic Protocol) Adult mouse Mouse dissection tools 70% ethanol spray solution 1.5-ml microcentrifuge tubes, sterile 1× PBS, sterile: diluted from 10× phosphate-buffered saline without calcium and magnesium (Sigma, cat. no. P7059) to 1× with water, and sterilized by passing through a 0.22-μm filter 70-μm cell strainer (BD Falcon; optional) 22-G needle and 1-ml syringe (for bone marrow progenitors) 23-G needle and 1-ml syringe (for fetal liver progenitors) 25-G needle and 1-ml syringe (for fetal liver progenitors) NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

Prepare TLX1 producer cells 1. Grow TLX1 retroviral producer cell to 90% to 100% confluence in a 100-mm tissue culture dish. Irradiate the cell monolayer with 3 Gy and return to the tissue culture incubator. Although irradiation of the producer cell line is recommended, it is not absolutely necessary since the immortalized hematopoietic cells grow in suspension and the producer cells are adherent. Therefore, any contaminating retroviral producer cells that get carried over after the coculture period can eventually be removed by serial passage of the populations on tissue culture plates. If gene-targeted cells containing a neo targeting construct are immortalized with the MSCV-HOX11 retrovirus, then any remaining producer cells can also be eliminated by selection in Geneticin. If the producer cell line is not irradiated, the plating density should be slightly lower (e.g., ∼50% to 70% confluent in a 100-mm tissue culture dish); otherwise, producer cells may become confluent and lift off the dish.

Prepare hematopoietic progenitor cells

TLX1 Immortalization of Hematopoietic Progenitors

For bone marrow–derived hematopoietic progenitors 2a. Euthanize an adult mouse by carbon dioxide asphyxiation using compressed gas, or according to institutionally approved methods, and immediately spray the fur over the legs with 70% ethanol to disinfect. Dissect out both femurs and cut off ends. Methods of euthanasia other than carbon dioxide asphyxiation may be used; however, we do not know if they would affect the cells.

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3a. Hold the cut femur with forceps over a sterile 15-ml conical centrifuge tube and use a 1-ml syringe fitted with a 22-G needle to flush the bone marrow space with 1 ml IMDMES-15 with glutamine and pen/strep (0.5 ml in each direction). Repeat to obtain a total of 2 ml of cell suspension, pooling the sample. 4a. Repeat step 3a, using the other femur, and combine the samples (total 4 ml of cell suspension). 5a. Cap and vortex the tube. Allow debris to settle by gravity at room temperature for 5 min. 6a. Transfer the supernatant to fresh tube and discard the pellet, which contains mostly bone fragments and fibrous debris. Alternatively, bone marrow fragments and debris can be removed from the flushed marrow by filtering through a sterile 70-μm cell strainer (BD Falcon).

7a. Measure the viable cell concentration using a hemacytometer (UNIT 1C.3). 8a. Centrifuge ∼3 × 106 cells 5 min at ∼200 × g, 4◦ C, in a tabletop tissue culture centrifuge. 9a. Aspirate and discard the supernatant, and resuspend the cell pellet in 10 ml of coculture medium.

For fetal liver–derived progenitors 2b. Euthanize a pregnant female mouse at embryonic day 13.5 or 14.5 (E13.5 to E14.5) by carbon dioxide asphyxiation using compressed gas, or according to institutionally approved methods. Methods of euthanasia other than carbon dioxide asphyxiation may be used; however, we do not know if they would affect the cells.

3b. Immediately spray the fur over the abdomen with 70% ethanol. Using forceps and sharp scissors (first soaked in 70% ethanol and wiped dry), incise the abdominal wall and extract the embryos. Place the embryos in a sterile 100-mm petri dish containing sterile 1× PBS, and separate them by cutting with scissors. 4b. Open the yolk sacs and remove the fetuses, keeping them in the 1× PBS. Using blunt dissection with sterile forceps and scissors, remove the fetal livers from the embryos’ abdomens. The liver should come out easily at this stage of development.

5b. Place one fetal liver per sterile 1.5-ml microcentrifuge tube containing 1 ml IMDMES-15 with glutamine and pen/strep. 6b. Prepare a single-cell suspension by drawing the tissue through a 23-G needle five times, followed by a 25-G needle five times. 7b. Pool cell suspensions from several fetal livers into a 15-ml conical centrifuge tube. Determine viable cell concentration using a hemacytometer (UNIT 1C.3). 8b. Centrifuge ∼3 × 106 cells 5 min at ∼200 × g, 4◦ C, in a tabletop tissue culture centrifuge. 9b. Aspirate and discard the supernatant, and resuspend the cell pellet in 10 ml coculture medium. Specific progenitor cell populations from bone marrow or fetal liver can also be immunophenotypically isolated by fluorescence-activated cell sorting (FACS) and used for coculture with TLX1-expressing retroviral producer cell lines. For markers that can be

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used to select the progenitors at the various stages of differentiation, see Akashi et al. (2000) and Pronk et al. (2007).

Immortalize and passage the EB cells 10. Wash the irradiated producer cell monolayer (from step 1) once with 5 ml IMDMES-15 with glutamine and pen/strep. 11. Gently add the bone marrow or fetal liver cell suspension (∼3 × 105 cells/ml) to the irradiated cell monolayer. Incubate to coculture for 2 to 3 days, without changing the medium. 12. At the end of the coculture period, gently agitate the dishes to remove suspension cells loosely adherent to the monolayer and transfer the supernatant to a 15-ml conical tube. 13. Centrifuge 5 min at ∼200 × g, 4◦ C. 14. Resuspend the cell pellet in 3 ml of immortalized cell medium and incubate in a 60-mm tissue culture dish. Over the next few days, clusters of proliferating cells in suspension should be visible.

15. Select for TLX1-expressing cells (step 16) as early as 48 to 72 hr following the coculture, or passage cells 1:10 to 1:20 every 3 to 4 days, with gentle pipetting up and down to disrupt cell clusters. In our experience, TLX1 immortalized cells are almost always IL-3 dependent for their growth and survival. Therefore, it is critical to include IL-3 in the medium after the coculture and all subsequent steps. However, the IL-3-containing medium can be supplemented with other cytokines/growth factors, e.g., SCF, Epo (Keller et al., 1998; Yu et al., 2002; Riz et al., 2007).

Select for TLX1-expressing immortalized cells 16a. For cells transduced with the neo-carrying retrovirus cell line (GP+E-86/MSCVHOX11): Add Geneticin to a final concentration of 400 μg/ml. 16b. For cells transduced with the hyg-carrying retrovirus cell line (GP+E86/MSCVhyg-HOX11): Add hygromycin B to a final concentration of 400 μg/ml. While selection for retrovirally transduced cells with Geneticin or hygromycin B is recommended, it is not strictly required since TLX1-expressing cells will exhibit a growth advantage over nontransduced cells during in vitro passaging (Yu et al., 2002; Su et al., 2006) and nontransduced cells should eventually disappear from the cultures due to senescence. However, this may take as long as several months, especially for long-lived primary cells, e.g., mast cells.

17. Clone Geneticin- or hygromycin B-resistant cells in methylcellulose as described in Support Protocol 2 or by limiting dilution in liquid culture (see Fuller et al., 2001) using immortalized cell medium containing the appropriate selective antibiotic. The IL-3 containing methylcellulose and medium can be supplemented with other cytokines/growth factors such as SCF, Epo, etc. (Keller et al., 1998; Yu et al., 2002; Riz et al., 2007).

18. Freeze clonal cell lines as described in Support Protocol 3. TLX1 Immortalization of Hematopoietic Progenitors

19. Characterize the clonal lines, depending on the particular application, e.g., by identifying the hematopoietic cell lineage(s) represented, and their relative state of maturation (Support Protocol 4).

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MAINTENANCE OF TLX1 (HOX11) RETROVIRAL PRODUCER CELL LINES

SUPPORT PROTOCOL 1

The GP+E-86/MSCV-HOX11 producer cell line is a murine fibroblast cell line that exports helper-free replication-defective recombinant retrovirus carrying the TLX1 (HOX11) cDNA and the neo drug resistance gene (Hawley et al., 1994b). The GP+E86/MSCVhyg-HOX11 producer cell line is a murine fibroblast cell line that exports helper-free replication-defective recombinant retrovirus carrying for the TLX1 (HOX11) cDNA and the hyg drug resistance gene (Hawley et al., 1994b). The cell lines are available from the authors upon request. Maintain both cell lines in Dulbecco’s modified Eagle medium with 4.5 g/liter glucose + 10% (v/v) heat-inactivated calf serum or fetal bovine serum (FBS). (It is acceptable to use calf serum for maintaining the producer cells to reduce costs, but FBS should be used in the Basic Protocol and Alternate Protocol.) Add Geneticin or hygromycin B to a final concentration of 400 μg/ml or 200 μg/ml, respectively, to maintain selection for the retroviral constructs. Passage cells by tryspinization with 0.05% trypsin/0.53 mM EDTA at room temperature. As soon as the cells loosen from the plate, neutralize the trypsin by adding serum to a final concentration of 10%. Split the cells at a dilution of 1:7 to 1:10 every 2 to 3 days, when they are typically 90% to 100% confluent. The following patents have been issued that relate to the establishment of TLX1 (HOX11)immortalized ES cell–derived lines and use of the cells:

United States Patent No. 5,874,301 Title: Embryonic Cell Populations and Methods to Isolate Such Populations Inventors: Keller, G.M., Hawley, R.G., and Choi, K. Assignee: National Jewish Medical and Research Center, Denver, Colo. Issued: February 23, 1999 United States Patent No. 6,110,739 Title: Method to Produce Novel Embryonic Cell Populations Inventors: Keller, G.M., Hawley, R.G., and Choi, K. Assignee: National Jewish Medical and Research Center, Denver, Colo. Issued: August 29, 2000 United States Patent No. 6,555,318 Title: Method for Identification of Cell Growth or Differentiation Factors Inventors: Keller, G.M., Hawley, R.G., and Choi, K. Assignee: National Jewish Medical and Research Center, Denver, Colo. Issued: April 29, 2003 United States Patent No. 6,576,433 Title: Method for Identification of Cell Growth or Differentiation Factors Inventors: Keller, G.M., Hawley, R.G., and Choi, K Assignee: National Jewish Medical and Research Center, Denver, Colo. Issued: June 10, 2003 United States Patent No. 7,374,934 Title: Cell Populations and Methods of Production Thereof Inventors: Keller, G.M., Hawley, R.G., and Choi, K. Assignee: National Jewish Medical and Research Center, Denver, Colo. Issued: May 20, 2008. Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 2

CLONING OF IMMORTALIZED CELLS IN METHYLCELLULOSE This protocol describes a method for cloning Geneticin- or hygromycin B-resistant immortalized cells in methylcellulose-based medium. Alternatively, the cells can be cloned by limiting dilution (see Fuller et al., 2001) in immortalized cell medium containing the appropriate selective antibiotic, depending on which TLX1 (HOX11) retroviral producer cell line was used for the immortalization (Geneticin for MSCV-HOX11 and hygromycin for MSCVhyg-HOX11).

Materials Suspension of immortalized cells (Basic Protocol or Alternate Protocol) Methylcellulose medium (see recipe) Immortalized cell medium (see recipe), containing the appropriate selection antibiotic Tissue culture centrifuge (refrigerated, benchtop centrifuge with swinging bucket rotor) 15-ml centrifuge tubes (sterile) 5-ml syringe fitted with a 16-G needle 35-mm tissue culture dish (Falcon, cat. no. 35-1008) 150-mm bacterial dishes (Falcon, cat. no. 35-1058) Inverted microscope 2- to 20-μl micropipettor and sterile, disposable tips 96-well flat bottom tissue culture plate (Costar, cat. no. 3596) 24-well tissue culture plate (Falcon, cat. no. 35-3047) Additional reagents and equipment for counting cells (UNIT IC.3) 1. Pipet the suspension of immortalized cells up and down several times to break up clumps. Determine the cell concentration using a hemacytometer (UNIT IC.3). 2. Centrifuge appropriate volumes of cell suspension containing 200, 2000, and 20,000 cells separately in 15-ml centrifuge tubes for 5 min at 200 × g, 4◦ C. 3. Carefully aspirate off the supernatant, leaving a small amount (∼50 μl) of medium behind so as not to disturb the cell pellet. 4. Add 3.5 ml methylcellulose mix containing the appropriate selection antibiotic (Geneticin or hygromycin) to each tube, using a 5-ml syringe fitted with a 16-G needle. Carefully mix the cell-methylcellulose mixture by drawing it up and down in the syringe several times. Try to avoid making air bubbles. 5. Using the syringe and needle, add ∼1.5 ml of each of the mixtures into separate 35-mm dishes. Place six dishes in a 150-mm bacterial dish, including an extra uncovered 35-mm dish filled with sterile water at the center (to prevent drying out of the methylcellulose). Cover the 150-mm bacterial dish, and place it in the tissue culture incubator. 6. Examine the cultures daily. Typically, colonies are large enough to pick after ∼1 week.

7. Choose dishes with well separated colonies. Under the inverted microscope, carefully aspirate selected colonies using a pipettor fitted with a disposable, sterile tip, and set at ∼10 μl. TLX1 Immortalization of Hematopoietic Progenitors

8. Immediately add the aspirated colony to a well of a 96-well tissue culture plate containing 150 μl immortalized cell medium with the appropriate selection antibiotic. Wash out the tip several times by aspirating and expelling the liquid medium in the

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well. Be careful not to cross-contaminate neighboring wells when passing the pipettor over the plate. Most of the colonies should grow nicely in the liquid culture.

9. Examine the cultures daily, and expand them to 1-ml cultures in 24-well plates once they are growing well. 10. Expand to progressively larger culture volumes as necessary, and/or freeze aliquots as described in Support Protocol 3.

CRYOPRESERVATION OF IMMORTALIZED CELLS The following protocol describes a method we have found effective for cryopreservation of immortalized cells.

SUPPORT PROTOCOL 3

Materials Suspension of immortalized cells (Basic Protocol, Alternate Protocol, or Support Protocol 2) Immortalized cell medium (see recipe), ice cold 2× freezing medium (see recipe), ice cold Tissue culture centrifuge (refrigerated, benchtop centrifuge with swinging bucket rotor) 15-ml centrifuge tubes (sterile) 1.8-ml cryovials (Nunc, cat. no. 377267) −80◦ C freezer Liquid nitrogen storage tank 1. Centrifuge 1 × 107 immortalized cells in 15-ml centrifuge tubes 5 min at 200 × g, 4◦ C. 2. Aspirate and discard the supernatant, and resuspend the cells in 5 ml ice-cold immortalized cell medium. 3. Add 5 ml ice-cold 2× freezing medium. Pipet up and down to mix, and dispense 1-ml aliquots into each of 10 cryovials (∼1 × 106 cells/vial). 4. Cap vials and place on ice until they can be placed into a −80◦ C freezer. 5. Incubate vials in a −80◦ C freezer for 1 to 2 days, and then transfer to a liquid nitrogen tank for long-term storage. This procedure can be scaled up to freeze more cells. If there is poor viability after thawing, one can try substituting heat-inactivated fetal bovine serum (FBS) for the immortalized cell medium in step 2.

IMMUNOPHENOTYPIC ANALYSIS OF IMMORTALIZED CELL LINES BY FACS

SUPPORT PROTOCOL 4

Characterization of the clonal lines will depend on the particular application. However, identification of the hematopoietic cell lineages represented by the cell line, and their relative state of maturation, is common. A basic protocol for immunophenotypic analysis of hematopoietic lineage assignment by flow cytometry (FACS) and a list of suggested antibodies (Table 1F.7.1) are provided below. Additional procedures may include histochemical staining of the cells and/or reverse transcriptase-polymerase chain reaction (RT-PCR), northern blot analysis, or western blot analysis for lineage-specific and maturation specific gene expression. For a more detailed description and protocols pertaining to hematopoietic lineage analysis, see Baron (2005).

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Table 1F.7.1 Specific Cell-Surface Markers and Antibodies for Different Murine Hematopoietic Lineages

Lineage

Marker

Recommended supplier and antibody clone

Erythroid

CD71 (early) Ter119 (late)

BD Pharmingen clone C2 BD Pharmingen clone Ter-119

Megakaryocytic

CD41a (early) CD61 CD42b (late)

BD Pharmingen clone MSReg30 BD Pharmingen clone 2C9.G2 Emfret clone Xia.G5

Granulocytic

Gr-1 (Ly-6G)

BD Pharmingen clone RB6-8C5

Macrophage

Mac-1

BD Pharmingen clone M1/70

B lymphocyte

CD19 B220

BD Pharmingen clone 1D3 BD Pharmingen clone RA3-6B2

T lymphocyte

CD3

BD Pharmingen clone 17A2

a CD41 is also present on earlier multipotential progenitor cells.

Materials Cell line suspensions (Basic Protocol or Alternate Protocol) FACS wash buffer (see recipe) Fluorochrome-conjugated antibodies and isotype matched control antibodies (see Table 1F.7.1); choice of fluorochromes depends on the flow cytometer being used and the combination of markers being analyzed Tissue culture centrifuge (refrigerated, benchtop centrifuge with swinging bucket rotor) 15-ml conical centrifuge tube 12 × 75–mm FACS tubes (BD Falcon, cat. no. 35-2054) Flow cytometer (FACS analyzer) Additional reagents and equipment for counting cells (UNIT IC.3) 1. Determine cell concentration of suspension using hemacytometer (UNIT IC.3). 2. Centrifuge 2 × 106 cells in a 15-ml conical centrifuge tube 5 min at 200 × g, 4◦ C. 3. Aspirate and discard the supernatant. Wash the cells two times with 1 ml FACS wash buffer, gently vortexing or pipetting up and down to mix. Centrifuge 5 min at 200 × g, 4◦ C, between washes. 4. After the last wash, resuspend the cells with 100 μl FACS wash buffer. 5. Add 0.2 μg fluorochrome-conjugated antibody (typically, 1 μl of the recommended antibodies listed in Table 1F.7.1) or equivalent isotype-matched antibody to an identical aliquot of control cells. Incubate 30 min at 4◦ C, in the dark, with occasional agitation of the tube to mix the cells. The amount of antibody suggested is a starting point. Titrations may be needed to optimize cell staining.

6. Centrifuge the cells 5 min at 200 × g, 4◦ C. 7. Aspirate and discard the supernatant. Wash the cells two times with 1 ml FACS wash buffer, centrifuging 5 min at 200 × g, 4◦ C, between washes. TLX1 Immortalization of Hematopoietic Progenitors

8. Resuspend the final cell pellet with 1 ml of FACS wash buffer and transfer to 12 × 75–mm FACS tubes. Place samples on ice and keep in the dark. The stained cells can be stored for several hours at 4◦ C before analyzing.

9. Analyze on a flow cytometer (FACS analyzer).

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Ascorbic acid stock solution, 5 mg/ml Prepare a stock solution of 5 mg L-ascorbic acid (Sigma; A-4544)/ml autoclaved water and sterilize by passing through a 0.22-μm filter. Prepare ascorbic acid stock solution fresh for each differentiation procedure.

Coculture medium Fetal bovine serum, ES grade (FBS-ES; Hyclone), heat-inactivated (15%, v/v, final) 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final 200 mM glutamine (2 mM final) Interleukin-3 (IL-3; R & D Systems; 10 ng/ml final) Interleukin-6 (IL-6; R & D Systems; 2 ng/ml final) Interleukin-11 (IL-11; R & D Systems; 5 ng/ml final) Stem cell factor (SCF; R & D Systems; 50 ng/ml final) Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 1 month at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C.

FACS wash buffer 10× phosphate-buffered saline, without calcium and magnesium (CMF-PB; Sigma, cat. no. P7059) diluted to 1× (final) with water Fetal bovine serum (FBS; Hyclone), heat-inactivated (1%, v/v, final) Sodium azide (0.1%, w/v, final) Store up to 6 months at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C.

Freezing medium, 2× Fetal bovine serum (FBS; Hyclone), heat-inactivated (60%, v/v, final) Dimethyl sulfoxide (DMSO; Sigma, cat. no. D8418; 20%, v/v, final) Immortalized cell medium (see recipe; 20%, v/v, final) Store indefinitely at −20◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C. The addition of DMSO to the solution is an exothermic reaction and will produce heat. Cool the 2× freezing medium on ice before adding to cells for cryopreservation.

Gelatin 0.1% (w/v) Dilute 10× phosphate-buffered saline without calcium and magnesium (CMF-PBS; Sigma, cat. no. P7059), to 1× with water. Sterilize by passing through a 0.22-μm filter. Add 0.5 g gelatin (Sigma, cat. no. G-1890) to 500 ml of the 1× CMF-PBS and autoclave. Store up to 1 month at 4◦ C, in the absence of microbial contamination.

Gelatinized flasks Prior to addition of ES cells, add enough 0.1% gelatin solution (see recipe) to coat the surface of the flask. Incubate at room temperature for 20 min. Remove excess gelatin solution before adding cells. Store up to 6 months at room temperature.

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IMDM-ES-5 with MTG and pen/strep Fetal bovine serum, ES grade (FBS-ES; Hyclone), heat-inactivated (5%, v/v, final) α-monothioglycerol (MTG; Sigma, cat. no. M-1753; 12.4 μl/liter final) 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 1 month at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C. Use MTG stock from a bottle that has been open for <6 months.

IMDM-ES-15 with glutamine and pen/strep Fetal bovine serum, ES grade (FBS-ES; Hyclone), heat-inactivated (15%, v/v, final) 2 mM L-glutamine 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 2 months at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C.

IMDM-ES-15 with MTG, LIF, and pen/strep Fetal bovine serum, ES grade (FBS-ES; Hyclone), heat-inactivated (15%, v/v, final) α-monothioglycerol (MTG; Sigma, cat. no. M-1753; 12.4 μl/liter final; first diluted from stock, in IMDM, if necessary) Leukemia inhibitory factor (LIF; R & D Systems; 1000 U/ml final) 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 1 month at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C. Use MTG stock from a bottle that has been open for <6 months.

Immortalized cell medium Fetal bovine serum (FBS; standard tissue culture grade; Hyclone; heat-inactivated; 15%, v/v, final) 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final L-glutamine (2 mM final) Recombinant murine interleukin-3 (IL-3; R & D Systems, cat. no. 403-ML; 10 ng/ml final) or X630-recombinant IL-3-conditioned medium (Karasuyama and Melchers, 1988; 10%, v/v, final) Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 2 months at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C.

Methylcellulose medium TLX1 Immortalization of Hematopoietic Progenitors

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2.5× Methocult stock solution (StemCell Technologies, cat. no. 03234) diluted to 1× (final) with immortalized cell medium (see recipe) L-glutamine (2 mM final) continued

Current Protocols in Stem Cell Biology

100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final IL-3 (R & D systems, cat. no. 403-ML; 10 ng/ml final) Geneticin (Invitrogen, cat. no. 10131-035; 400 mg/ml final) or hygromycin B (400 mg/ml final) for selection (depending on the TLX1 retroviral producer cell line used for immortalization) Prepare fresh for each experiment Prewarming the methylcellulose at 37◦ C for a few minutes in a water bath makes it easier to work with.

Primary ES differentiation medium 1:1 fetal bovine serum (FBS; Hyclone), heat-inactivated/plasma-derived serum (PDS; Antech), 15% (v/v) final L-glutamine (2 mM final) α-monothioglycerol (MTG; 0.004%, v/v, final; first diluted from stock, in IMDM, if necessary) 5 mg/ml ascorbic acid stock solution (see recipe; 50 μg/ml final) Protein-free hybridoma medium (PFHMII; Invitrogen, cat. no. 23600-026; 5%, v/v, final) 100× penicillin/streptomycin solution (Sigma; P4333): 1× (100 U penicillin/ml; 100 μg streptomycin/ml) final IL-11 (5 ng/ml final) Recombinant murine or rat stem cell factor (rSCF; also called KL, kit-ligand; R & D Systems; 50 ng/ml final) Isocove’s modified Dulbecco’s medium (IMDM; Invitrogen, cat. no. 12440) to make up the final volume Store up to 1 month at 4◦ C Heat inactivation of FBS is carried out for 30 min at 56◦ C.

COMMENTARY Background Information The diverged homeobox-containing gene TLX1 (formerly called HOX11) was first identified in T cell acute lymphoblastic leukemia carrying the t(10;14)(q24;q11) chromosomal translocation (Dube et al., 1991; Hatano et al., 1991; Kennedy et al., 1991; Lu et al., 1991). In this translocation, the T cell receptor δ chain gene regulatory sequences become juxtaposed to the 5 promoter region of the TLX1 gene, leading to aberrant TLX1 expression in the T cell lineage. A similar translocation, t(7;10)(q35;q24), places the T cell receptor β chain gene regulatory sequences into the TLX1 gene 5 promoter region, causing a similar effect. Subsequent studies showed that retroviral expression of TLX1 immortalizes murine hematopoietic cells derived from bone marrow cells, fetal liver, yolk sac, and in vitro differentiated ES cells at high frequency (Hawley et al., 1994a; Keller et al., 1998; Yu et al., 2002; Zhang et al., 2002; Yu et al., 2003; Owens et al., 2006). A wide variety of hematopoietic cell types have been immortalized, including

erythroid, megakaryocytic, myelocytic, and monocytic lineages. Interestingly, there are no reports of TLX1 immortalization of T lymphocyte progenitors, even though its misexpression disrupts T lymphocyte differentiation and leads to T cell leukemia in humans (Owens et al., 2006). Essentially all of the reported TLX1-immortalized cell lines are IL-3 dependent for their growth and survival. The mechanism of TLX1 cell immortalization remains incompletely understood. An intact homeodomain is required, suggesting direct transcriptional effects due to DNA binding activity (Owens et al., 2003). However, overexpression of TLX1 deregulates genes involved in controlling G1/S cell cycle progression and disrupts a G2/M cell-cycle checkpoint, apparently by indirect mechanisms mediated in part via inhibition of PP1/PP2A phosphatase activity (Kawabe et al., 1997; Riz and Hawley, 2005). TLX1 expression also alters the subcellular distribution of CREBbinding protein (CBP), a transcriptional coactivator with histone acetyltransferase activity

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(Riz et al., 2007). It seems likely that multiple mechanisms contribute to TLX1 immortalization of hematopoietic progenitor cells. Examples of how TLX1 immortalized cell lines have been utilized experimentally include: (1) structure-function study of the GATA transcriptional cofactor Friend of GATA-1 (FOG-1; Cantor et al., 2002); (2) investigation of the role of the SHP-2 tyrosine phosphatase in interleukin-3 signaling (Yu et al., 2003); (3) chromatin immunoprecipitation assays demonstrating FOG-1 facilitation of GATA-1 chromatin occupancy (Pal et al., 2004); (4) establishment of a link between the proapoptotic BCL-2 family member Bid and the DNA-damage response (Zinkel et al., 2005); (5) testing the function of mitoferrin, a novel mitochondrial iron transporter (Shaw et al., 2006); and (6) transcription factor target gene identification using Cre-mediated excision in TLX1 immortalized cells generated from SCLflox/flox (H. Mikkola, pers. commun.) and Runx-1flox/flox mice (M. Yu and A.B. Cantor, unpub. observ.). On a note of caution, the process of immortalization by TLX1 results in perturbed or arrested differentiation potential (Hawley et al., 1994a; Keller et al., 1998; Dixon et al., 2007; Riz et al., 2007). This should be taken into consideration when designing cell fate experiments utilizing the TLX1 immortalized cell lines.

Critical Parameters and Troubleshooting

TLX1 Immortalization of Hematopoietic Progenitors

In our experience, the protocols for TLX1 immortalization of hematopoietic progenitors described in this unit yield cell lines quite readily. When using in vitro differentiated ES cells as a source of target cells, the time of EB harvest is an important consideration. We typically use EBs harvested at 7 days of culture. We have generated cell lines from 6-day-old EBs, but the efficiency seems to be lower. The stage of EB differentiation used for the immortalization will also theoretically affect whether primitive or definitive hematopoietic progenitors are targeted. We reported TLX1 immortalization of embryonic precursors with both primitive and definitive hematopoietic potential from day 7 EBs (Keller et al., 1998). Two of the lines had the capacity to generate cells with both primitive and definitive erythroid potential. A third line was limited to definitive erythroid potential, but also had myeloid potential. We have also found that the quality and reproducibility of the ES cell in vitro differentiation declines when using older stock solu-

tions of MTG. We therefore try to avoid using MTG stocks that are greater than 6 months old. Most cells will continue to replicate after the coculture with TLX1 retroviral producer cells regardless of whether they are transduced by the TLX1 retroviruses. However, these nontransduced cells should eventually undergo senescence. Mast cells are particularly longlived and may be retained in primary cultures with IL-3 for many weeks before naturally senescing. Although one can “wait out” the eventual senescence of these nonimmortalized cells by serial passaging, one can also select for the TLX1-immortalized cells earlier by including Geneticin or hygromycin B (depending on the TLX1 retroviral vector used) in the medium beginning 48 hr after coculture with the retroviral producer cells. If not used up front, resistance to Geneticin or hygromycin B should be tested to ensure that the cells have been transduced with the retroviral vectors. In addition, immunoblot analysis of the final cell lines should be performed to ensure expression of TLX1. This migrates as an ∼37-kDa protein and can be detected using affinity-purified anti-TLX1 polyclonal antibody (HOX11 C18; Santa Cruz Biotechnology). Because immortalized hematopoietic cells typically grow in suspension, it is relatively easy to separate them from any residual retroviral producer cells, which are adherent; this is also easily accomplished by cloning in methylcellulose. However, the monolayer of producer cells can be irradiated with 3 Gy prior to adding the hematopoietic target cells to ensure that the producer cells do not contaminate future cell lines. Finally, MSCV-HOX11 retroviral producer cells can be eliminated by selection in Geneticin if the immortalized cell lines are derived from gene-targeted cells expressing the neo gene. In our experience, TLX1-immortalized cells are strictly dependent on IL-3 for survival and growth. Therefore, it is critical to add IL-3 to the medium during the coculture with the retroviral producer cells, or at least soon after. We have used conditioned media from WEHI-3B cells (Lee et al., 1982) or X630-recombinant IL-3 cells (Karasuyama and Melchers, 1988) as a source of IL-3, but have found better cell growth and viability using recombinant murine IL-3 (10 ng/ml, R&D Systems).

Anticipated Results After the 3-day coculture period, proliferation of the transferred suspension cells should be apparent within a few days, with cells

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growing as clusters in liquid medium. Even after cloning the cell lines, the wild-type cell cultures may remain heterogeneous with a mixture of self-renewing multipotent progenitor cells of various potentials, and differentiated progeny at various stages of maturation. Use of gene-targeted cells as the source of immortalized cells may lead to more homogenous cell populations if the disrupted gene is required for a specific stage of hematopoietic differentiation. For example, we generated a FOG-1−/− cell line from in vitro differentiated FOG-1−/− ES cells (Cantor et al., 2002; Fig. 1F.7.1). This cell line is blocked in erythroid and megakaryocytic maturation (as expected), and therefore has a more homogenous morphology. Although genetic complementation of the cells by retrovirally expressed FOG1 rescues the terminal maturation block, only about 35% of the cells differentiate in this case, as measured by staining with o-dianisidine for hemoglobin production. A large proportion of cells remain undifferentiated, presumably reflecting the self-renewing population of immortalized progenitor cells.

Time Considerations Overall, generation of TLX1-immortalized cell lines can be accomplished within a couple of weeks. For the in vitro differentiation protocol, the ES cells are first differentiated for 7 days. The resultant EBs are then dissociated and directly cocultured on prepared TLX1 retroviral producer cell monolayers for 3 days. At this point, the transduced cells can be cultured as a pool, or cloned immediately in the presence of Geneticin or hygromycin B (and IL-3). Colonies should be detectable within about 2 weeks and expanded accordingly. Cell lines should be maintained by splitting at a dilution of ∼1:10-1:20 every 3 to 4 days. The doubling time of the cells may be in the range of ∼36 to 48 hr.

Acknowledgments A.B.C would like to thank Stuart Orkin and Catherine Porcher for advice with development of the TLX1 immortalization of in vitro differentiated ES cells. A.B.C. is supported by National Institutes of Health (NIH) grants R01HL075705 and R01HL82952). R.G.H. is supported by NIH grants R01HL65519 and R01HL66305) and by an Elaine H. Snyder Cancer Research Award and a King Fahd Endowed Professorship from the George Washington University Medical Center.

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Owens, B.M., Zhu, Y.X., Suen, T.C., Wang, P.X., Greenblatt, J.F., Goss, P.E., and Hawley, R.G. 2003. Specific homeodomain-DNA interactions are required for HOX11-mediated transformation. Blood 101:4966-4974. Owens, B.M., Hawley, T.S., Spain, L.M., Kerkel, K.A., and Hawley, R.G. 2006. TLX1/HOX11mediated disruption of primary thymocyte differentiation prior to the CD4+ CD8+ doublepositive stage. Br. J. Haematol. 132:216-229. Pal, S., Cantor, A.B., Johnson, K.D., Moran, T.B., Boyer, M.E., Orkin, S.H., and Bresnick, E.H. 2004. Coregulator-dependent facilitation of chromatin occupancy by GATA-1. Proc. Natl. Acad. Sci. U.S.A. 101:980-985.

Hawley, R.G., Fong, A.Z., Lu, M., and Hawley, T.S. 1994a. The HOX11 homeobox-containing gene of human leukemia immortalizes murine hematopoietic precursors. Oncogene 9:1-12.

Perkins, A.C. and Cory, S. 1993. Conditional immortalization of mouse myelomonocytic, megakaryocytic and mast cell progenitors by the Hox-2.4 homeobox gene. EMBO J. 12:38353846.

Hawley, R.G., Lieu, F.H., Fong, A.Z., and Hawley, T.S. 1994b. Versatile retroviral vectors for potential use in gene therapy. Gene Ther. 1:136138.

Pinto do, O.P., Richter, K., and Carlsson, L. 2002. Hematopoietic progenitor/stem cells immortalized by Lhx2 generate functional hematopoietic cells in vivo. Blood 99:3939-3946.

Kamps, M.P. and Wright, D.D. 1994. Oncoprotein E2A-Pbx1 immortalizes a myeloid progenitor in primary marrow cultures without abrogating its factor-dependence. Oncogene 9:3159-3166.

Pronk, C.J., Rossi, D.J., Mansson, R., Attema, J.L., Norddahl, G.L., Chan, C.K., Sigvardsson, M., Weissman, I.L., and Bryder, D. 2007. Elucidation of the phenotypic, functional, and molecular topography of a myeloerythroid progenitor cell hierarchy. Cell Stem Cell 1:428-442.

Karasuyama, H. and Melchers, F. 1988. Establishment of mouse cell lines which constitutively secrete large quantities of interleukin 2, 3, 4 or 5, using modified cDNA expression vectors. Eur. J. Immunol. 18:97-104. Kawabe, T., Muslin, A.J., and Korsmeyer, S.J. 1997. HOX11 interacts with protein phosphatases PP2A and PP1 and disrupts a G2/M cell-cycle checkpoint. Nature 385:454-458. Keller, G., Wall, C., Fong, A.Z., Hawley, T.S., and Hawley, R.G. 1998. Overexpression of HOX11 leads to the immortalization of embryonic precursors with both primitive and definitive hematopoietic potential. Blood 92:877887. Kennedy, M.A., Gonzalez-Sarmiento, R., Kees, U.R., Lampert, F., Dear, N., Boehm, T., and Rabbitts, T.H. 1991. HOX11, a homeoboxcontaining T-cell oncogene on human chromosome 10q24. Proc. Natl. Acad. Sci. U.S.A. 88:8900-8904. Lavau, C., Szilvassy, S.J., Slany, R., and Cleary, M.L. 1997. Immortalization and leukemic transformation of a myelomonocytic precursor by retrovirally transduced HRX-ENL. EMBO J. 16:4226-4237.

TLX1 Immortalization of Hematopoietic Progenitors

Lu, M., Gong, Z.Y., Shen, W.F., and Ho, A. 1991. The tcl-3 proto-oncogene altered by chromosomal translocation in T-cell leukemia codes for a homeobox protein. EMBO J. 10:2905-2910.

Lee, J.C., Hapel, A.J., and Ihle, J.N. 1982. Constitutive production of a unique lymphokine (IL 3) by the WEHI-3 cell line. J. Immunol. 128:23932398.

Riz, I. and Hawley, R.G. 2005. G1/S transcriptional networks modulated by the HOX11/TLX1 oncogene of T-cell acute lymphoblastic leukemia. Oncogene 24:5561-5575. Riz, I., Akimov, S.S., Eaker, S.S., Baxter, K.K., Lee, H.J., Marino-Ramirez, L., Landsman, D., Hawley, T.S., and Hawley, R.G. 2007. TLX1/HOX11-induced hematopoietic differentiation blockade. Oncogene 26:4115-4123. Shaw, G.C., Cope, J.J., Li, L., Corson, K., Hersey, C., Ackermann, G.E., Gwynn, B., Lambert, A.J., Wingert, R.A., Traver, D., Trede, N.S., Barut, B.A., Zhou, Y., Minet, E., Donovan, A., Brownlie, A., Balzan, R., Weiss, M.J., Peters, L.L., Kaplan, J., Zon, L.I., and Paw, B.H. 2006. Mitoferrin is essential for erythroid iron assimilation. Nature 440:96-100. Schnabel, C.A., Jacobs, Y., and Cleary, M.L. 2000. HoxA9-mediated immortalization of myeloid progenitors requires functional interactions with TALE cofactors Pbx and Meis. Oncogene 19:608-616. Su, X., Drabkin, H., Clappier, E., Morgado, E., Busson, M., Romana, S., Soulier, J., Berger, R., Bernard, O.A., and Lavau, C. 2006. Transforming potential of the T-cell acute lymphoblastic leukemia-associated homeobox genes HOXA13, TLX1, and TLX3. Genes Chromosomes Cancer 45:846-855.

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Suh, HC., Leeanansaksiri, W., Ji, M., Klarmann, K.D., Renn, K., Gooya, J., Smith, D., McNiece, I., Lugthart, S., Valk, P.J., Delwel, R., and Keller, J.R. 2008. Id1 immortalizes hematopoietic progenitors in vitro and promotes a myeloproliferative disease in vivo. Oncogene 27:5612-5623. Templin, C., Kotlarz, D., Rathinam, C., Rudolph, C., Schatzlein, S., Ramireddy, K., Rudolph, K. L., Schlegelberger, B., Klein, C., and Drexler, H. 2008. Establishment of immortalized multipotent hematopoietic progenitor cell lines by retroviral-mediated gene transfer of betacatenin. Exp. Hematol. 36:204-215. Varnum-Finney, B., Xu, L., Brashem-Stein, C., Nourigat, C., Flowers, D., Bakkour, S., Pear, W.S., and Bernstein, I.D. 2000. Pluripotent, cytokine-dependent, hematopoietic stem cells are immortalized by constitutive Notch1 signaling. Nat. Med. 6:1278-1281.

Yu, W.M., Hawley, T.S., Hawley, R.G., and Qu, C.K. 2003. Catalytic-dependent and -independent roles of SHP-2 tyrosine phosphatase in interleukin-3 signaling. Oncogene 22:5995-6004. Zhang, P., Nelson, E., Radomska, H.S., IwasakiArai, J., Akashi, K., Friedman, A.D., and Tenen, D.G. 2002. Induction of granulocytic differentiation by 2 pathways. Blood 99:4406-4412. Zhang, X.B., Schwartz, J.L., Humphries, R.K., and Kiem, H.P. 2007. Effects of HOXB4 overexpression on ex vivo expansion and immortalization of hematopoietic cells from different species. Stem Cells 25:2074-2081. Zinkel, S.S., Hurov, K.E., Ong, C., Abtahi, F.M., Gross, A., and Korsmeyer, S.J. 2005. A role for proapoptotic BID in the DNA-damage response. Cell 122:579-591.

Yu, W.M., Hawley, T.S., Hawley, R.G., and Qu, C.K. 2002. Immortalization of yolk sac-derived precursor cells. Blood 100:3828-3831.

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Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts from Human Embryonic Stem Cells

UNIT 1F.8

Maria Elena Stavropoulos,1 Isabella Mengarelli,1 and Tiziano Barberi1 1

Beckman Research Institute of City of Hope, Duarte, California

ABSTRACT This unit describes a protocol for the derivation of multipotent mesenchymal precursors from human embryonic stem cells (hESCs). hESCs cultured at low density in the presence of a chemically defined serum-free medium are induced to adopt an endomesodermal fate and later a mesenchymal phenotype. FACS sorting for the surface antigen CD73 is used to purify mesenchymal precursors able to differentiate into fat, bone, cartilage, and skeletal muscle cells. Enrichment in mesenchymal precursors with a myogenic potential is achieved via an additional FACS sorting for the embryonic skeletal muscle surface C 2009 by John Wiley marker N-CAM. Curr. Protoc. Stem Cell Biol. 9:1F.8.1-1F.8.10.  & Sons, Inc. Keywords: hESC r FACS r hESC-MP

INTRODUCTION This unit describes a two-step protocol for the derivation of multipotent hESC mesenchymal precursors (hESC-MP). hESCs cultured at low density in the presence of a chemically defined serum-free medium are induced to adopt an endomesodermal fate and later a mesenchymal phenotype. FACS sorting for the surface antigen CD73 is used to purify mesenchymal precursor cells able to differentiate into bone, cartilage, and skeletal muscle cells. An additional FACS sorting for the embryonic skeletal muscle surface marker N-CAM leads to enrichment in mesenchymal precursor cells with myogenic potential. First the induction of the differentiation of small hESC colonies seeded on murine embryonic fibroblasts (MEFs) or on fibronectin-coated plates into mesenchymal cells for 3 weeks, followed by an additional 1 week of cell expansion, is described. After the expansion phase, a FACS sorting is performed to purify mesenchymal progenitors and myogenic cells. See Figure 1F.8.1 for a schematic representation of the procedure.

DIFFERENTIATION OF hESCS INTO MESENCHYMAL PROGENITORS: ENDOMESODERMAL INDUCTION

BASIC PROTOCOL 1

The first step in this protocol is trypsinization of hESCs to a single-cell suspension (5 to 10 min at 37◦ C), which is then plated at a density of ∼1000 cells/cm2 on MEFs. Alternatively, small clumps of hESCs can be plated on feeder-free fibronectin-coated plates. The survival of single hESCs plated in feeder-free conditions is very low; therefore, small hESC clumps or colonies (consisting of ∼50 cells) need to be plated on this substrate. With continued culture and appropriate medium changes the colonies will begin to differentiate, becoming mesenchymal progenitors, hESC-MP. Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1F.8.1-1F.8.10 Published online June 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f08s9 C 2009 John Wiley & Sons, Inc. Copyright 

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Figure 1F.8.1 Schematic representation of the procedure. Time course and details are given for the two sequential steps leading to the isolation of hESC-MPs (see Basic Protocol 1), and the enrichment in myogenic hESC-MPs (see Alternate Protocol 1). Details are also given for the terminal differentiation of hESC-MPs and myogenic hESC-MPs derivatives (see Basic Protocol 2 and Alternate Protocol 2).

Materials hESCs hESC medium (see recipe) ITS medium (see recipe) 60-mm 1 μg/ml fibronectin-coated culture dishes (Biocoat; BD Falcon) Phosphate-buffered saline, Ca2+ /Mg2+ -free (CMF-PBS; Cellgro, Mediatech) 0.05% (w/v) trypsin/EDTA (Invitrogen) MEM medium (see recipe) FACS sorting buffer (see recipe) Anti-human CD73 monoclonal antibody, PE conjugated (BD Pharmingen) or PE-conjugated IgG isotype control (BD Pharmingen) Gelatin (0.1% in water; Specialty Media, Millipore) Aspirator 37◦ C incubator 15-ml centrifuge tubes Tissue culture grade dishes (60-mm; BD Falcon) Hemacytometer MoFlo FACS sorter (Dako Cytomation) Prepare hESCs 1. After replating hESCs, maintain cells under standard hESC conditions in 3 ml hESC medium for 3 to 4 days to allow colony formation. Change the medium every day. 2. When colonies reach a size of 30 to 50 cells, switch the medium to 3 ml ITS medium (day 0). Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts

3. Replace the medium with 3 ml fresh ITS medium every 2 to 3 days (also see Fig. 1F.8.2).

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hESC day 0

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Figure 1F.8.2 hESC cultured in ITS medium show a developmental progression toward endomesodermal fate and later mesenchyme.

4. At day 14, replate cells on 60-mm fibronectin-coated dishes at a split ratio of 1:3. To replate cells, remove ITS medium, wash with 2 ml PBS, aspirate PBS, and add 0.6 ml of 0.05% trypsin/EDTA and incubate 5 min in a 37◦ C incubator. 5. Inactivate trypsin with 1 ml MEM medium, collect cells in a 15-ml tube, and centrifuge 5 min at 960 × g, room temperature. 6. Aspirate medium and resuspend pellet in 3 ml ITS medium, plate 1 ml cell suspension per 60-mm tissue culture dish and add 3 ml of ITS medium to each dish. 7. At day 21 switch medium to 3 ml MEM medium and maintain the culture for an additional 7 days. 8. Replace medium with 3 ml fresh MEM medium every 2 to 3 days for duration of 7 days. 9. If necessary, (if cells are confluent), split cells again at a ratio 1:3 following steps 4 to 6 but using MEM medium. The cells are differentiated when their morphology appears to be different than the ES cells.

Isolate hESC-MP cells by FACS sorting 10. Aspirate the medium from differentiated hESC cultures and wash each dish two times with 2 ml PBS. 11. Add 0.6 ml of 0.05% trypsin/EDTA and incubate 5 min in a 37◦ C incubator. 12. Inactivate trypsin by adding 1 ml MEM medium, collect cells in a 15-ml tube, and centrifuge 5 min at 960 × g, room temperature.

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Figure 1F.8.3 Representative FACS sorting for the isolation of the CD73+ percursors. A shows the isotype control. B shows CD73 PE. Reproduced with permission from Barberi et al. (2007).

13. Aspirate medium, resuspend pellet in 1 ml FACS sorting buffer, and count cells using a hemacytometer chamber (UNIT 1C.3). 14. Resuspend cells in FACS sorting buffer to a concentration of 106 cells/ml and stain with the appropriate amount of anti-CD73 PE antibody or PE-conjugated isotype IgG (10 μl/106 cells, as recommended by manufacturer). 15. Incubate cells 30 min on ice in the dark. 16. Add 10 ml PBS to the cell suspension and centrifuge 5 min at 960 × g, room temperature. 17. Resuspend pellet in the appropriate volume of FACS sorting buffer, as recommended by manufacturer, and proceed with FACS sorting (see also Fig. 1F.8.3). 18. Collect CD73-positive cells (hESC-MP) and replate them in an appropriately sized gelatinized tissue culture dish in the presence of MEM medium at a cell density of ∼5 × 103 cells/cm2 . 19. Expand the cells by spitting 1:3 when cultures reach 70% confluency (about every other day). The hESC-MPs can now either be expanded for further differentiation experiments (see Support Protocol) or cryopreserved in modified MEM medium (35% FBS) with 10% DMSO. SUPPORT PROTOCOL

Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts

MAINTENANCE OF hESC CULTURES Undifferentiated hESCs, H1 (WA-01, XY, passages 40 to 60) and H9 (WA-09, XX, passages 40 to 60) cell lines, are passaged on mitotically inactivated MEF (Specialty Media, Millipore). The hESCs are maintained under the growth conditions and passaging techniques described previously (Zhang et al., 2001).

Materials hESC cultures, undifferentiated and grown on MEFs Phosphate buffered saline (PBS; Cellgro, Mediatech) Collagenase IV/dispase (see recipe)

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hESC medium (see recipe) Mitotically inactivated MEFs (Specialty Media, Millipore) MEF medium (see recipe) Gelatin (0.1% in water; Specialty Media, Millipore) 37◦ C incubator 15-ml tubes Aspirator Tissue culture grade dishes (60-mm; BD Falcon) 1. When cultures of hESCs reach colony sizes (∼1 mm), aspirate the medium, wash with 3 ml PBS, and remove PBS. 2. Add 2 ml collagenase IV/dispase and incubate 40 min at 37◦ C. 3. Transfer the hESC clumps to a 15-ml tube and add 10 ml of hESC medium and gently pipet up and down. 4. Centrifuge cells 5 min at 680 × g, room temperature. 5. Aspirate medium and resuspend pellet in 12 ml hESC medium. 6. Replate the small clumps of hESCs at a split ratio of 1:4 on fresh mitotically inactivated MEFs preseeded the day before on gelatin-coated dishes at a cell density of 2 × 104 cells/cm2 in MEF medium. Remove the MEF medium before adding the hESCs.

ENRICHMENT FOR hESC-MP WITH MYOGENIC POTENTIAL BY FACS SORTING

ALTERNATE PROTOCOL 1

For the enrichment of hESC-MPs with myogenic potential, FACS sorting for the embryonic skeletal muscle marker N-CAM is performed on expanded, purified (CD73+ ) hESC-MPs using a MoFlo FACS sorter (see Fig. 1F.8.4). Myogenic hESC-MPs can be expanded in culture and need to be passaged before they reach 70% confluency. Under these conditions, they retain their myogenic potential for more than 12 to 15 passages.

Materials hESC-MP (<70% confluent dish) Phosphate buffered saline (PBS; Cellgro, Mediatech) 0.05% (w/v) trypsin/EDTA (Invitrogen) MEM medium (see recipe) FACS sorting buffer (see recipe) Anti-N-CAM monoclonal antibody (clone 5.1H11; DSHB) Fluorochrome-conjugated secondary antibody (chosen by investigator) 0.1% (w/v) gelatin (Embryomax, Millipore) Aspirator 37◦ C incubator 15-ml tubes Hemacytometer MoFlo FACS sorter (Dako Cytomation) Tissue culture dishes (various sizes) 1. Aspirate medium from a <70% confluent dish of hESC-MPs and briefly wash two times with 2 ml PBS. 2. Add 0.6 ml of 0.05% trypsin/EDTA and incubate 5 min at 37◦ C. 3. Inactivate trypsin by adding 1 ml MEM medium, collect cells in a 15-ml tube, and centrifuge 5 min at 960 × g, room temperature. Current Protocols in Stem Cell Biology

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Figure 1F.8.4 Representative FACS sorting for the enrichment of skeletal muscle cells from hESC-MPs based on N-CAM immunoreactivity. Reproduced with permission from Barberi et al. (2007).

4. Aspirate medium and resuspend pellet in 1 ml FACS sorting buffer. Count cells using a hemacytometer (UNIT 1C.3). 5. Resuspend cells in FACS sorting buffer to a concentration of 106 cells/ml and stain with the appropriate amount of anti N-CAM antibody (10 μl/106 cells). 6. Incubate 30 min on ice. 7. Add 10 ml PBS to the cell suspension and centrifuge 5 min at 960 × g, room temperature. 8. Resuspend pellet in FACS sorting buffer, as recommended by manufacturer, and stain with the appropriate fluorochrome-conjugated secondary antibody. Also stain control cells (not previously stained with N-CAM) with the secondary antibody. 9. Incubate 30 min on ice in the dark. 10. Add 10 ml PBS to the cell suspension and centrifuge 5 min at 960 × g, room temperature. 11. Resuspend the pellet in the appropriate volume of FACS sorting buffer and proceed with FACS sorting (see also Fig. 1F.8.3). 12. Collect N-CAM-bright positive cells (myogenic hESC-MP) and replate at 2.5 × 103 cells/cm2 in the appropriately sized gelatinized tissue culture dish in the presence of MEM medium. Myogenic hESC-MPs can also be cryopreserved in modified MEM medium (35% FBS) with 10% DMSO. BASIC PROTOCOL 2

TERMINAL SKELETAL MUSCLE DIFFERENTIATION OF MYOGENIC hESC-MP Myogenic hESC-MP can be terminally differentiated into mature myocytes able to fuse and form myotubes.

Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts

Materials Myogenic hESC-MP cultures (see Alternate Protocol 1) MEM medium (see recipe) N2 medium (see recipe)

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Figure 1F.8.5 Example of skeletal myocytes differentiated from myogenic hESC-MPs. DAPI staining (A), sarcomeric myosin (MF-20) in green, and Myogenin in red (B).

1. Grow myogenic hESC-MP in MEM medium until they become 100% confluent. Cells are fed every other day and it takes 2 to 3 days to reach 100% confluency.

2. Aspirate MEM medium and replace with the same volume of N2 medium. 3. Replace medium with fresh N2 medium every 2 days. 4. Grow the cultures until they reach the desired stage of differentiation. Mature myocytes become visible after 24 hr of growth in N2 medium and fusion of mature myocytes into myotubes starts at day 4 in N2 medium (also see Fig. 1F.8.5).

TERMINAL DIFFERENTIATION OF MESENCHYMAL DERIVATIVES This section briefly describes protocols (not originally set up by the authors’ laboratory) for the terminal differentiation of hESC-MP that are extensively described elsewhere (Pittenger et al., 1999; Barberi et al., 2005; Barberi et al., 2007; see also Fig. 1F.8.6).

ALTERNATE PROTOCOL 2

Adipocytic differentiation Adipocytic differentiation of hESC-MPs (see Basic Protocol 1) is achieved on 100% confluent cells by exposure for 2 to 4 weeks to 1 mM dexamethasone, 10 μg/ml insulin, and 0.5 mM isobutylxanthine (all reagents available from Sigma) in MEM medium. Cells are fed with fresh medium containing the additives every 2 to 3 days. No cell passage is required. At the end of the treatment, 50% to 70% of the cells will have fat granules, indicating adipocytic differentiation. Fat granules can be visualized by Oil-Red-O staining. Chondrocytic differentiation Chondrocytic differentiation of hESC-MPs (see Basic Protocol 1) is achieved using the pellet culture system (5 × 105 cells/pellet) and exposure for 3 to 4 weeks to 5 ng/ml transforming growth factor β3 (TGFβ3; R&D Systems) and 200 M ascorbic acid (AA; Sigma) in MEM medium. Alcian blue staining is performed to detect cartilage-specific extracellular matrix. Cells are fed with fresh medium containing the additives every 2 to 3 days. Osteogenic differentiation Osteogenic differentiation is achieved by culturing hESC-MPs (see Basic Protocol 1) at low density (1–2.5 × 103 cells/cm2 ) on tissue culture–treated dishes by exposure to

Embryonic and Extraembryonic Stem Cells

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A

B

C

Oil-Red-O

Alcian Blue

ALP/DAPI

fat

cartilage

bone

Figure 1F.8.6 Representative differentiation of hESC-MPs into adipocytes (A), condrocytes (B), and osteocytes (C). Reproduced with permission from Barberi et al. (2007).

10 mM β-glycero-phosphate (Sigma), 1 μM dexamethasone, and 200 μM AA in MEM medium for 3 to 4 weeks. Cells are fed with fresh medium containing the additives every 2 to 3 days. No cell passage is required. Positive Von Kossa reaction or Alizarin Red staining predict calcium deposition (ossification).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Collagenase/dispase solution 170,000 U/mg collagenase IV (Invitrogen) 1.7 U/mg dispase (Invitrogen) To prepare collagenase/dispase solution, prepare a 1:1 ratio with a concentration of 1 mg/ml in hESC medium (see recipe). Store up to 30 days at 4◦ C.

FACS sorting buffer CMF-PBS (Cellgro, Mediatech) containing: 2% (v/v) FBS (Invitrogen) EDTA solution 0.263 mM (Irvine Scientific) Store up to 6 months at 4◦ C hESC medium

Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts

DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (Invitrogen) 10 mM non-essential amino acids (Invitrogen) 1 mM L-glutamine (Invitrogen) 1× penicillin/streptomycin (from stock, e.g., Invitrogen) continued

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50 mM 2-mercaptoethanol (Invitrogen) 4 ng/ml bFGF (Invitrogen) Store up to 2 weeks at 4◦ C ITS medium DMEM/F12 (Invitrogen) containing: 1.55 g/liter D-glucose (Sigma) 2.438 g/liter NaHCO3 (Fisher Scientific) 5 mg/liter bovine insulin (Sigma) dissolved in 5 ml of 10 mM NaOH 50 mg/liter human apotransferrin (Sigma) 30 nM sodium selenite (Sigma) 1× penicillin/streptomycin (from stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C N2 medium DMEM/F12 (Invitrogen) containing: 1.55 g/liter D-glucose (Sigma) 2.0 g/liter NaHCO3 (Fisher Scientific)) 100 mg/liter human apotransferrin (Sigma) 25 mg/liter bovine insulin (Sigma) dissolved in 5 ml of 10 mM NaOH 100 uM putrescine (Sigma) 20 nM progesterone (Sigma) 30 nM sodium selenite (Sigma) 1× penicillin/streptomycin (from stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C MEF medium High-glucose DMEM (Invitrogen) containing: 10% (v/v) FBS (Invitrogen) 2 mM L-glutamine 1× penicillin/streptomycin (from stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C MEM medium Alpha MEM (Invitrogen) containing: 10% (v/v) heat-inactivated FBS (Invitrogen) 2 mM L-glutamine 1× penicillin/streptomycin (from stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C COMMENTARY Background Information hESCs are pluripotent cells deriving from fertilized eggs at the blastocyst stage. Because of their early embryonic origin, they can be induced to recapitulate in vitro the earliest stages of human development and can be differentiated into a variety of tissues and specialized cells. Therefore, there is a lot of interest in harnessing the full potential of hESC through the establishment of protocols for the directed differentiation into the cell type of interest. The two-step protocol in this unit allows the generation of a large number of expandable mes-

enchymal progenitors with osteogenic, chondrogenic, adipogenic, and myogenic potential. Although mesenchymal precursor cells can be isolated from a variety of adult tissue, hESCMPs offer the following advantages: extensive proliferative potential, no need for a donor, and more importantly, the ability to differentiate into skeletal muscle cells. In addition, other protocols for the isolation of mesenchymal precursors from hESCs are not based on FACS purification and their myogenic potential has not been assessed (Lian et al., 2007).

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Critical Parameters and Troubleshooting For an efficient generation of mesenchymal precursor cells, hESC colonies must be small in size before starting the ITS medium induction. This step will minimize the spontaneous neural differentiation that can also be achieved in the presence of ITS medium. For culture of hESCs on fibronectin-coated plates, seeding of a single cell suspension is not recommended, because of the poor survival of single hESCs. Seeding small hESC clusters (∼50 cells/cluster) is recommended, if this substrate is favored by the investigator. Although this protocol is very efficient in generating mesenchymal precursor cells and results are highly reproducible, the efficiency of obtaining myogenic hESC-MP can vary among hESC lines. In addition, investigators should avoid allowing the hESC-MP to become overly confluent during expansion to prevent loss of myogenic potential. Retention of full differentiation potential of hESC-MP in long-term expansion (>20 passages) needs to also be evaluated.

Anticipated Results This protocol generates large numbers of hESC-derived mesenchymal precursors (hESC-MPs) suitable for numerous applications, such as developmental studies, drug testing, and future cell therapy. FACS sorting– mediated purification yields multipotent mesenchymal precursors with osteogenic, chondrogenic, adipogenic, and myogenic potential. hESC-MPs can be expanded, in the appropri-

ate conditions, for more than 30 passages allowing the generation of up to 109 cells.

Time Consideration To generate and isolate mesenchymal precursor cells from undifferentiated hESCs, 4 weeks are required.

Acknowledgements The authors would like to thank Lorenz Studer (Memorial Sloan-Kettering Cancer Center) for his help and contribution in the development of this protocol.

Literature Cited Barberi, T., Willis, L.M., Socci, N.D., and Studer, L. 2005. Derivation of multipotent mesenchymal precursors from human embryonic stem cells. PLoS Med. 2:e161. Barberi, T., Bradbury, M., Dincer, Z., Panagiotakos, G., Socci, N.D., and Studer, L. 2007. Derivation of engraftable skeletal myoblasts from human embryonic stem cells. Nat. Med. 13:642-648. Lian, Q., Lye, E., Yeo, K.S., Tan, E.K.W., SaltoTellez, M., Liu, T.M., Palanisamy, N., Oakley, R.M.E., Lee, E.H., Lim, B., and Lim, S.K. 2007. Derivation of clinically compliant MSCs from CD105+, CD24-differentiated human ESCs. Stem Cells 25:425-436. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., and Marshak, D.R. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284:143147. Zhang, S.C., Wernig, M., Duncan, I.D., Brustle, O., and Thomson, J.A. 2001. In vitro differentiation of transplantable neural precursors from human embryonic stem cells. Nat. Biotech. 19:11291133.

Differentiation of Multipotent Mesenchymal Precursors and Skeletal Myoblasts

1F.8.10 Supplement 9

Current Protocols in Stem Cell Biology

Derivation of Vasculature from Embryonic Stem Cells

UNIT 1F.9

Jeffrey N. Lindquist,1,2 David A. Cheresh,2 and Evan Y. Snyder1 1 2

The Burnham Institute for Medical Research, La Jolla, California UCSD Moores Cancer Center, La Jolla, California

ABSTRACT The formation of the multicellular vascular system is critical to the growth, development, and viability of an organism, and many embryonic lethal mouse knockouts are due to vascular defects. Unfortunately, the complex nature, and many cell types involved in vasculogenesis and angiogenesis has stymied in vitro models of vascular formation. This unit describes a system that allows human embryonic stem cells to differentiate and spontaneously form vascular networks via both vasculogenesis and angiogenesis in the C 2010 context of the three germ layers. Curr. Protoc. Stem Cell Biol. 12:1F.9.1-1F.9.6.  by John Wiley & Sons, Inc. Keywords: endothelial cells r vascular patterning r differentiation r angiogenesis r embryoid body

INTRODUCTION This unit describes a protocol for the differentiation of pluripotent hESCs into vascular networks in the presence of all three developmental germ layers. Mechanical disruption of hESCs and the subsequent growth in suspension to form embryoid bodies (EBs) facilitates the formation of the germ layers. The seeding of EBs onto a two-dimensional matrix allows proliferation, migration, and patterning of many cell types, including vascular endothelial cells (ECs). These ECs undergo vasculogenesis, the formation of a vascular plexus by the aggregation of ECs, and angiogenesis, the formation of new vessels by sprouting from an existing vessel, to form a vascular plexus that undergoes maturation. This unit will begin with hESC dissection and EB formation, followed by EB differentiation and analysis. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

hESC DISSECTION, EMBRYOID BODY FORMATION, AND SEEDING hESCs are routinely passaged by mechanical dissection, and the same protocol is performed for EB formation (Oh et al., 2005).

BASIC PROTOCOL

Materials hESCs, starting from macroscopic colonies (<1-mm diameter) grown on MEFs or human fibroblast feeders hESC growth medium (see recipe) Enzymatically passaged hESC Current Protocols in Stem Cell Biology 1F.9.1-1F.9.6 Published online March 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f09s12 C 2010 John Wiley & Sons, Inc. Copyright 

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Type I collagen (Sigma, cat. no. C3867) Phosphate-buffered saline (PBS; Invitrogen, cat. no. 14040-133) Differentiation medium (see recipe) Reagents for detecting endothelial markers (Table 1F.9.1) 6-well low-binding tissue culture plates (Corning, cat. no. 3471) 20- to 200-μl pipets and tips, sterile 12-well tissue culture–treated culture plates 37◦ C incubator 15- and 50-ml tubes Stereomicroscope (preferably with a heated stage) Plate hESC for embryoid body formation For embryoid body formation from hESC colony fragments: 1a. Aspirate hESC growth medium and add 1 ml of fresh hESC growth medium without bFGF to each well of a 6-well plate. 2a. Scrape around the periphery of a hESC colony to remove the feeders directly contacting the colony. Dissect only round, opaque colonies that do not have the “cratered” or flattened differentiated phenotype.

3a. Using a sterile pipet tip or needle, score the colony in a grid and gently dislodge the pieces. 4a. Gently, remove all the medium and colony pieces and place into 1 well of a 6-well low-binding culture plate. This is day 0. 5a. Add 2 to 3 ml hESC growth medium without bFGF every other day to day 7. Remove medium if well is getting full, and gently disperse EBs from the center of the well when adding fresh medium to prevent them from clumping together. There may be some dead cells/debris in the well—this will not affect EB formation. For hESC embryoid body formation from single-cell suspension:

1b. To form embryoid bodies from a single-cell suspension (Ellerstrom, 2007), seed enzymatically passaged hESC in a low-binding culture dish at a density of 1 × 106 to 1 × 107 cells/ml in hESC growth medium without bFGF in a total volume of 4 ml/well in a 6-well plate. Proceed to step 6. The cells will spontaneously aggregate into EBs over several days. There will be some dead cells and debris visible, but this should not affect EB formation or viability.

Differentiate the embryoid bodies 6. On day 7, coat a 12-well tissue culture–treated plate with 10 μg/ml of type I collagen in PBS at 37◦ C for at least 1 hr. Coating overnight also works well. Table 1F.9.1 Endothelial Markers and Related Detection Reagents

Derivation of Vasculature from Embryonic Stem Cells

Endothelial marker

Antibody

Supplier

CD-31

Mouse-anti-CD31

Zymed, cat. no. 37-0700

VE-Cadherin

Goat-anti-VE-Cad

Santa Cruz, cat. no. SC-6458

Ulex Europa Lectin

Vector Laboratories, cat. no. RL1062

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day 0

day 4

day 7

day 11

day 14

day 17

day 21

A

B

C

Figure 1F.9.1 Time course of hES vascular differentiation. hES colonies (day 0) are dissociated and cultured in suspension to form EBs for 7 days. EBs attach to collagen-coated dish and proliferate and migrate outward until day 21. Staining with U. E. Lectin labels endothelial cells that spontaneously arise under these conditions. A cross-section of the culture illustrates the different regions that arise. The initiation area under the EB (A), an area undergoing endothelial and vessel maturation (B), and an area where angiogenic endothelial cells sprout and migrate towards the leading edge of the culture (C). Corresponding bright-field image is shown below the U. E. Lectin staining. Scale bar = 200 μm.

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7. Remove medium and EBs (they should be several hundred microns wide at this point) and place into a 15-ml tube and allow the EBs to pellet by gravity for 5 to 10 min. The EBs will sink to the bottom, but any debris in the medium will not.

8. Aspirate the medium above the EB clump, leaving 0.5 ml medium above the pellet. 9. Add 5 ml differentiation medium to the tube, and remove all EBs. Add EBs to a 50-ml tube with differentiation medium to a total volume of 48 ml. 10. Mix gently and seed 2 ml of EBs per well into the coated 12-well plate using a 25-ml pipet. Ensure reasonably balanced distribution of EBs/well by bubbling air up the pipet every 2 to 3 wells during seeding. Seed duplicate plates. If EBs are unevenly distributed, use a 5-ml pipet to combine a sparse well(s) and a dense well(s) and re-allocate more evenly.

11. Allow EBs to attach for 2 days, and then feed the EBS every other day by removing ∼1.5 ml and adding 2 ml fresh differentiation medium. Do not remove all medium during aspirations, and add fresh medium VERY SLOWLY. Culture until day 21. The EBs are prone to coming off the dish/tearing and should be handled very carefully.

Analyze differentiation 12. Fix the cultures and stain for endothelial markers (see Table 1F.9.1) at any time during the differentiation (see Fig. 1F.9.1 for the timeline). By day 10 of the differentiation, the EBs should be attached, with some cells spreading out. By days 14 to 16, endothelial clusters and primitive tubes are forming in the expanding “brim” of the sombrero. By day 21, the sombreros will have expansive, multi-layered “brims” containing endothelial vessel networks. The cultures are suitable for fixing with either 4% paraformaldehyde or methanol:acetone (1:1), and both of these fixations are compatible with immuno-fluorescence staining with antibodies. Check with your Environmental Health and Safety Department regarding disposal of these fixatives.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Differentiation medium DMEM (high glucose; Invitrogen, cat. no. 10313) with: 1× GlutaMAX (Invitrogen) 1× N2 Supplement (Invitrogen, cat. no. 17502048) 20 ng/ml bFGF (PeproTech, cat. no. 100-18B; see recipe) 50 ng/ml VEGF (PeproTech, cat. no. 100-20; see recipe) Store up to 6 months at 4◦ C without growth factors Add growth factors immediately prior to use hESC growth medium

Derivation of Vasculature from Embryonic Stem Cells

DMEM/F12 (Invitrogen, cat. no. 11320-033) with: 20% (v/v) Knockout Serum Replacement (Invitrogen, cat. no. 10828-28) 10 mM non-essential amino acids 2 mM L-glutamine 50 mM 2-mercaptoethanol Store up to 1 week at 4◦ C

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Recombinant human growth factors Resuspend lyophilized bFGF or VEGF (PeproTech) to a final concentration of 100 μg/ml in PBS containing 0.1% (w/v) BSA. Store up to 12 months at −80◦ C, do not freeze/thaw aliquots.

COMMENTARY Background Information Human embryonic stem cells (hESCs) are cells isolated from the inner cell mass of human blastocysts; these cells maintain pluripotency—the ability to differentiate into many cell types—during culture in vitro (Thomson et al., 1998). As such, they provide the ability to investigate cell fate decisions and cell-cell interactions during the formation of multi-cellular tissues and organs. They may also provide a ready source of particular cell types. ESCs have been differentiated into all cell lineages in the body, and many specialized cell types (e.g., beta cells, dopaminergic neurons, endothelial cells, smooth muscle cells; Krenning et al., 2007, 2008; Elkabetz et al., 2008; Meier et al., 2008; Obata and Kasuga, 2009). The efficiency of generating pure populations of specific cell types is quite low, however, and frequently requires sequential addition of growth factors or specific matrices that make analysis of the differentiation process difficult (Krenning et al., 2007; Lee et al., 2007; Mohan et al., 2008; Narita et al., 2008). Conversely, the use of laissez faire differentiation, where hESCs are allowed to differentiate spontaneously down any lineage, provides embryogenesis in a dish where many different cell types emerge simultaneously. This approach is described here and allows the study of intrinsic cell patterning and cell fate decisions in a developmental context. The conditions used and the addition of various growth factors can enhance the differentiation of particular cell types (e.g., VEGF for endothelial cells), but all three germ layers are present and contribute to the overall culture milieu. The spontaneous emergence of cell lineages and cell types adjacent to one another can enhance or prevent the neighboring cell differentiation along a particular lineage. Under these conditions, there is a spontaneous emergence of vascular networks that exhibit the characteristic developmental processes vasculogenesis and angiogenesis in vitro.

with high quality ES cells is critical. The density of EBs during suspension culture should not be so dense that you observe large aggregates of EBs fused together. The EBs tend to cluster in the center of the well, but gentle shaking daily keeps them from aggregating. During the differentiation, the medium can become acidic, especially as the differentiation progresses and more cells are present, so changing the medium becomes essential and can be done daily, if needed. Also, consider the type of readout for the experiment when seeding out EBs; for imaging, a few EBs per well provides the most possibilities for staining, but for RNA or protein isolation, seeding 15 to 30 EBs/well in a 6-well dish may be the most practical for ease of harvesting.

Anticipated Results Generally, there are regions of well-formed vascular networks after 18 to 21 days of culture. However, there can be a variation between different EBs, and within specific regions of an individual EB, regarding the formation of vasculature. Not every single EB will develop a vascular network, and there is variation between each network. However, analysis of multiple wells makes interpretation and analysis of the characteristics and dynamics of vascular networks possible. The strength of this protocol is that endothelial networks appear de novo from precursor cells and in direct contact with adjacent cells. This in vitro model of vascular formation provides the most complete representation of embryonic vascular formation in a cell culture environment. Many existing human endothelial cell lines have limited growth potential and lose endothelial characteristics after multiple passages. The hESC system, however, allows for genetic manipulation of the pluripotent hESCs, which can be grown indefinitely and differentiated into vascular networks as needed to observe endothelial phenotypes.

Time Considerations Critical Parameters and Troubleshooting The formation of vascular endothelium during spontaneous differentiation requires that the starting hES cells be pluripotent, so starting

This assay requires up to 21 days for completion, so proper planning of experiments prior to starting is paramount. By preparing embryoid bodies every time the hES cells are split one can ensure that fresh material for

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future experiments are always on hand. Furthermore, since the rough timeline of vascular stages is known—angiogenic clusters form on days 12 to 14, vascular plexus form on days 14 to 18, and vascular plexus remodeling occurs from days 18 to 21—treatments to perturb or enhance endothelial cell formation, maturation, or patterning can be added at particular times to avoid using excessive amounts of reagents and cellular toxicity. In general, expect that 21 to 24 days will be required to finish culturing the differentiations and perform analysis of the experiment, be it immunofluorescence, RNA/DNA analysis, or protein analysis.

Literature Cited Elkabetz, Y., Panagiotakos, G., Al Shamy, G., Socci, N.D., Tabar, V., and Studer, L. 2008. Human ES cell-derived neural rosettes reveal a functionally distinct early neural stem cell stage. Genes Dev. 22:152-165. Ellerstrom, C., Strehl, R., Noaksson, K., Hyllner, J., and Semb, H. 2007. Facilitated expansion of human embryonic stem cells by single-cell enzymatic dissociation. Stem Cells 25:16901696. Krenning, G., Dankers, P.Y., Jovanovic, D., van Luyn, M.J., and Harmsen, M.C. 2007. Efficient differentiation of CD14+ monocytic cells into endothelial cells on degradable biomaterials. Biomaterials 28:1470-1479. Krenning, G., Moonen, J.R., van Luyn, M.J., and Harmsen, M.C. 2008. Vascular smooth muscle cells for use in vascular tissue engineering obtained by endothelial-to-mesenchymal trans-

differentiation (EnMT) on collagen matrices. Biomaterials 29:3703-3711. Lee, G., Kim, H., Elkabetz, Y., Al Shamy, G.,. Panagiotakos, G., Barberi, T., Tabar, V., and Studer, L. 2007. Isolation and directed differentiation of neural crest stem cells derived from human embryonic stem cells. Nat. Biotechnol. 25:1468-1475. Meier, K., Lehr, C.M., and Daum, N. 2008. Differentiation potential of human pancreatic stem cells for epithelial- and endothelial-like cell types. Ann. Anat. 191:70-82. Mohan, N., Nair, P.D., and Tabata, Y. 2008. A 3D biodegradable protein-based matrix for cartilage tissue engineering and stem cell differentiation to cartilage. J. Mater. Sci. Mater. Med. 20:4960. Narita, Y., Yamawaki, A., Kagami, H., Ueda, M., and Ueda, Y. 2008. Effects of transforming growth factor-beta 1 and ascorbic acid on differentiation of human bone-marrow-derived mesenchymal stem cells into smooth muscle cell lineage. Cell Tissue Res. 333:449459. Obata, A. and Kasuga, T. 2009. Stimulation of human mesenchymal stem cells and osteoblasts activities in vitro on silicon-releasable scaffolds. J. Biomed. Mater. Res. 91:11-17. Oh, S.K., Kim, H.S., Park, Y.B., Seol, H.W., Kim, Y.Y., Cho, M.S., Ku, S.Y., Choi, Y.M., Kim, D.W., and Moon, S.Y. 2005. Methods for expansion of human embryonic stem cells. Stem Cells 23:605-609. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147.

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Current Protocols in Stem Cell Biology

Isolation and Functional Characterization of Pluripotent Stem Cell–Derived Cardiac Progenitor Cells

UNIT 1F.10

Xiaojing Huang1 and Sean M. Wu1,2 1

Cardiovascular Research Center, Division of Cardiology, Massachusetts General Hospital, Boston, Massachusetts 2 Harvard Stem Cell Institute, Cambridge, Massachusetts

ABSTRACT The use of transgenic markers in pluripotent stem cells allows the facile isolation of transient cell populations that appear at certain phases of embryonic development. Here, we describe a procedure for deriving cardiac progenitors from mouse pluripotent stem cells carrying a GFP reporter under the control of an Nkx2.5 enhancer sequence. The cells are propagated under standard conditions and are differentiated using the hanging-droplet method with medium optimized for commitment to the cardiac lineage. Cardiac progenitors are isolated from the differentiation culture using ßuorescence-activated cell sorting (FACS) and can be cultured further for functional characterization and experimentation. The protocols described here can be applied to both embryonic and induced pluripotent stem cells and can easily be adapted to transgenic lines carrying other cardiac cell lineage C 2010 by John Wiley & reporters. Curr. Protoc. Stem Cell Biol. 14:1F.10.1-1F.10.14.  Sons, Inc. Keywords: mouse embryonic stem cells r induced pluripotent stem cells r cardiac progenitor cells r in vitro differentiation

INTRODUCTION In recent years, there has been growing interest among basic and clinical scientists in harnessing the tremendous developmental potential of pluripotent stem cells for understanding fundamental biological processes and disease pathogenesis. These cells, derived either from pre-implantation stage embryos of human or animal origin [embryonic stem (ES) cells], from neonatal or adult testis [mature adult germline stem cells (maGSC)], or by molecular reprogramming of differentiated somatic cells [induced pluripotent stem cell (iPS) cells], are believed to share similar transcriptional proÞles and molecular phenotypes in culture. The ability of these cells to differentiate in vitro into all lineages of the three germ layers has allowed the isolation and phenotypic characterization of various populations of progenitor and transient-amplifying cells prior to their terminal differentiation. It is believed that by identifying and isolating lineage-speciÞc stem/progenitor cells from pluripotent stem cells, one may then be able to translate the developmental potential of these cells into therapeutic applications for a wide range of neurological, cardiovascular, and hematological diseases. The objective of this unit is to describe, in sufÞcient detail, our methods for identifying and isolating cardiac progenitor cells from in vitro–differentiated mouse ES and iPS cells. It is anticipated that all readers with standard training in cell and molecular biology will be able to accomplish this by following the steps described here. Before we describe our basic protocol for isolating cardiac progenitor cells from pluripotent stem cells, it is worth mentioning the rationale and advantages for using Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1F.10.1-1F.10.14 Published online September 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01f10s14 C John Wiley & Sons, Inc. Copyright 

1F.10.1 Supplement 14

in vitro–differentiated ES or iPS cells rather than isolating them directly from developing embryos. First, working with early stage embryos can be technically difÞcult and costly, and the number of cells obtained may be insufÞcient for further experimentation. Pluripotent stem cells, on the other hand, can be expanded in large numbers to satisfy the cell number required for most studies. Second, the rapid doubling time of pluripotent stem cells allows for a short turnaround time between experiments. Third, pluripotent stem cells can be genetically modiÞed to carry lineagespeciÞc reporters and/or mutations, which would allow for the in vitro isolation of speciÞc cell types of interest that recapitulate disease phenotypes without incurring the time and cost associated with generating genetically modiÞed animals. These advantages, coupled with the potential uses of pluripotent stem cells in cell-based therapies, account for the growing popularity of these cells in regenerative biology and medicine. In this unit, we describe the isolation and characterization of mouse ES and iPS cellderived Nkx2.5+ cardiac progenitor cells. While a number of cardiac stem/progenitor cell populations from mouse and human ES cells have been described (Kattman et al., 2006; Moretti et al., 2006; Yang et al., 2008; Bu et al., 2009; Domian et al., 2009), the biological relationships between these cell populations remain to be clariÞed. Moreover, the protocols used by each laboratory to purify cardiac progenitor cells vary signiÞcantly. For example, the use of Flk1 to isolate multipotent cardiovascular progenitor cells relies on the addition of a cocktail of growth factors during differentiation (Kattman et al., 2006). When mouse ES cells are differentiated in the presence of serum and the absence of growth factor cocktail, very few cardiomyogenic Flk1+ cells can be isolated (S.M. Wu, unpub. observ.). Our approach here takes advantage of the availability of a committed cardiac progenitor cell marker, Nkx2.5, which has been well described to mark the Þrst identiÞable heart-forming cells in the developing embryo (Lints et al., 1993; Lyons et al., 1995; Stanley et al., 2002; Wu et al., 2006), as well as a transient postnatal population whose signiÞcance is still under investigation (Wu et al., 2006). Furthermore, we have shown that mouse ES cell–derived Nkx2.5+ cells can give rise speciÞcally to cardiomyocytes and smooth muscle cells both in vivo and in vitro (Wu et al., 2006). Although the protocol described here is based on studies using a transgenic Nkx2.5-eGFP ßuorescence reporter to identify cardiac progenitor cells, the methods for cell culture, differentiation, and progenitor cell isolation can be applied to other ES and iPS cells carrying appropriate cardiac lineage or surface markers. The Þrst protocol describes the general culturing and in vitro differentiation of mouse ES cells by the hanging-droplet method. Subsequent protocols describe the isolation, continued culture, and characterization of the FACS-puriÞed eGFP+ cardiac progenitor cells from Nkx2.5-eGFP ES cells. NOTE: This unit assumes that the user has some experience with the growth and maintenance of undifferentiated mouse embryonic stem (ES) cells; for an example protocol, see UNIT 1C.4. NOTE: The following procedures are performed in a Class II biological hazard ßow hood, using sterile equipment and solutions and proper aseptic technique. Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

NOTE: All incubations are performed in a humidiÞed 37◦ C, 5% CO2 incubator unless otherwise noted. NOTE: All centrifugations are done at room temperature in an Eppendorf Model 5810R benchtop centrifuge using an A-4-81 rotor.

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CULTURING AND IN VITRO DIFFERENTIATION OF PLURIPOTENT STEM CELLS

BASIC PROTOCOL 1

This protocol is adapted from the well-described hanging-droplet method of ES differentiation (Samuelson and Metzger, 2006; also see UNIT 1D.2), with slight modiÞcations designed to enhance cardiac lineage differentiation (Wobus et al., 1991, Takahashi et al., 2003). Fetal bovine serum is used to provide the differentiation stimulus, in contrast to other methods that rely on growth factor–supplemented serum-free medium, some of which are incapable of forming hanging droplets due to inadequate surface tension. This protocol can be adapted to generate large quantities of cardiac progenitor cells by scaling up accordingly. For high-throughput screening assays, a monolayer differentiation protocol using lineage-labeled ES or iPS cells in a 96-well plate format can be used (see the Alternate Protocol). Additionally, a hybrid protocol that incorporates both methods may be used for other applications. In such a protocol, cells differentiated in hanging-droplet cultures are dissociated at the desired time point and replated into 6-, 12-, 24-, or 96-well plates for further treatments and characterization (see Basic Protocol 2).

Materials Nkx2.5-eGFP transgenic ES or iPS cells passaged fewer than 30× since original derivation, in standard culture in 6-well plates on a feeder layer of growth-arrested mouse embryonic Þbroblasts (MEFs; UNIT 1C.3); MEFs should be seeded at 400,000 cells/well DMEM-ES medium (see recipe) 0.25% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25200-056) IMDM-ES medium (see recipe) 0.1% (w/v) gelatin solution (see recipe) Differentiation medium (see recipe) 10-cm cell culture dish 10- to 100-μl multichannel pipettor 150 × 15–mm slippable monoplate petri dishes (Fisher, cat. no. 08-757-14) Additional reagents and equipment for counting cells (UNIT 1C.3) Deplete MEFs from cell culture (2 days) 1. Culture ES or iPS cells in DMEM-ES medium in one well of a 6-well feeder layer plate. When cells reach 50% to 70% conßuency, aspirate the medium and add 0.5 ml of 0.25% trypsin/EDTA to each well of cells. Incubate the cells for 5 min, or until cells are in a single-cell suspension, as visualized under a microscope. 2. Add 2 ml of IMDM-ES medium to each well to inactivate the trypsin. 3. Centrifuge the cells 3 min at 200 × g, room temperature. Aspirate the supernatant and resuspend the cell pellet in ∼1 ml IMDM-ES medium. 4. Add 10 ml of IMDM-ES medium to a 10-cm cell culture dish precoated with 0.1% gelatin. 5. Add resuspended ES cells to the plate and pipet gently to redistribute cells evenly throughout the dish. Incubate for 2 days. During this time, the ES or iPS cells will continue to propagate (up to 1.5 × 107 cells in the 10-cm dish), while the number of growth-arrested MEFs remains constant at the 400,000 carried over from the from the single well; consequently, the Þnal number of MEFs is <3% of the total cell count. Embryonic and Extraembryonic Stem Cells

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Figure 1F.10.1 Embryoid body formation from hanging droplet. (A) Pipetting 11-μl droplets of cells onto the bottom of a 15-cm petri dish. (B) The dish is inverted to generate hanging droplets. The droplet density shown here (∼250 droplets/plate) yields the maximum number of EBs per plate. (C) A representative area of beating eGFP+ cells within an EB.

Form embryoid bodies (EBs) 6. At the end of 2 days, dissociate cells from the 10-cm dish using 2 ml of 0.25% trypsin/EDTA. Incubate for 5 min or until cells have lifted from the dish and inactivate the trypsin with 8 ml of differentiation medium. 7. Centrifuge cells 3 min at 200 × g, room temperature. Aspirate the supernatant and resuspend the cell pellet in ∼1 ml differentiation medium. 8. Count the live cells (UNIT 1C.3). Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

9. Dilute the cell suspension in differentiation medium to a Þnal concentration of 200,000 cells/ml. 10. Using a multichannel pipettor set at 11 μl (to compensate for ∼1 μl loss with each pipettor action), pipet ∼250 droplets onto the bottom of an uncoated 15-cm petri dish (Fig. 1F.10.1A).

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The number of cells from a 10-cm IMDM-ES culture (∼1.5 × 107 ) is sufÞcient for ∼25 of these 15-cm dishes. The droplets are pipetted onto the bottom of the dish rather than on the lid, as is typically done in other hanging-droplet methods, because in 2 days, the culture will be converted to a suspension in which the EBs will be free to attach to the dish for further expansion and development.

11. Replace the lid of each dish, and turn the dish upside-down. The droplets will hang from the bottom of the plate (Fig. 1F.10.1B). This conÞguration allows the cells to aggregate and form embryoid bodies (EBs). The day that the droplets are made is counted as day 0, with the subsequent differentiation days (e.g., day 1) counted from this time point.

12. Incubate the cells as hanging droplets for 2 days. On day 2, turn the dishes rightside-up and add 10 ml of differentiation medium to each 15-cm petri dish. Swirl to ensure even coverage of the dish. The EBs should be barely visible as small white specks ßoating in suspension. A wellformed EB should be spherically shaped and have a smooth, bright border, as viewed under a light microscope.

Differentiate the EBs 13. Monitor the cultures over the next 3 to 5 days. If the ES cells have successfully differentiated, a signiÞcant portion (>50%) of the EBs will adhere to the bottom of the dish while continuing to enlarge. The Þrst GFP+ cells can be detected as early as day 5; these represent the earliest cardiac progenitors. Beating clusters containing GFP+ cells (Fig. 1F.10.1C) will become visible under a ßuorescence microscope starting at day 7. With proper re-feeding of the culture plates (see step 14), these clusters can continue to beat and express GFP for the next 8 to 10 days. At desired time points during the course of differentiation, EBs may be isolated for ßow cytometric analysis, ßuorescence-activated cell sorting (FACS), immunocytochemistry, gene expression analysis, and other assays (see Basic Protocol 2). Differentiating cells secrete cytokines, growth factors, and other yet poorly deÞned chemical signals into the culture medium that are essential for proper differentiation. Therefore, the culture medium should not be removed before the cells have become committed to their respective lineages (usually by day 7). However, for continued EB growth and development beyond this time point, the medium will need to be replenished every 2 to 3 days.

14. At day 7, remove as much of the existing medium as possible through aspiration, taking care to not aspirate any unattached EBs. Add back 10 ml of fresh differentiation medium. For optimal results, repeat every 2 to 3 days.

IN VITRO DIFFERENTIATION OF PLURIPOTENT STEM CELLS IN A TWO-DIMENSIONAL HIGH-THROUGHPUT FORMAT

ALTERNATE PROTOCOL

For experiments involving chemical- or siRNA-based high-throughput screening of compounds or genes that regulate cardiac differentiation of ES or iPS cells, a monolayer differentiation assay in 96-well plate can be employed (Takahashi et al., 2003). If a ßuorescence reporter is used to quantify cardiac differentiation, a number of automated high-throughput detection systems, including ßow-cytometers, ßuorescent microscopes, and ßuorescent plate-readers, can be used.

Additional Materials (also see Basic Protocol 1) ES or iPS cells with MEFs depleted (see Basic Protocol 1) Collagenase/DNase solution (see recipe) HEPES-buffered saline (HBS; see recipe), optional

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Flow cytometry buffer (see recipe) Propidium iodide solution (1 mg/ml; Invitrogen, cat. no. P3566) 96-well cell culture plates, gelatin-coated Polypropylene cluster tubes for ßow cytometry (Corning Costar, cat. no. 4411) 1. Prepare ES or IPS cells for differentiation by depleting the MEF layer as described in Basic Protocol 1, steps 1 to 4. Because fewer cells are needed in general for monolayer differentiation compared with EB differentiation, a smaller size cell culture dish (<10 cm) may be used for the MEF depletion step.

2. Trypsinize the cells at the end of 2 days, centrifuge, and resuspend in ∼1 ml differentiation medium (see Basic Protocol 1, steps 6 and 7). 3. Count the live cells (UNIT 1C.3). 4. Add differentiation medium to bring the cells to 40,000 cells/ml. 5. Using a multichannel pipet, pipet 4000 cells (100 μl) into each well of a gelatincoated 96-well plate. The day during which this step takes place is counted as day 0, equivalent to day 0 of the EB differentiation protocol. Because the differentiating cells require signals that they secrete into the medium as they differentiate, do not exchange the medium prior to day 6, even if it becomes very acidic.

6. For ßow cytometry analysis prior to day 7, dissociate the differentiating cells using 50 μl 0.25% trypsin/EDTA per well for 5 min at 37◦ C. At day 7 or later, it will be necessary to use 50 μl/well collagenase/DNase solution for 1 hr at 37◦ C. 7. After digestion is complete, inactivate the enzyme by adding differentiation medium (trypsin) or HBS (collagenase), at a volume of at least 4× the volume of enzyme added. Centrifuge the cells 3 min at 200 × g, room temperature, using a centrifuge plate-holder, then aspirate the supernatant, taking care to avoid aspirating the cell pellets. 8. Add 100 μl of ßow cytometry buffer supplemented with 0.2% (v/v) propidium iodide solution to each well and pipet gently to resuspend cells. If a high-throughput ßow cytometer is not available, transfer the cell suspensions to a 96-well format rack of cluster tubes and add additional ßow cytometry buffer to each tube as needed to meet the minimum sample volume required by the machine. 9. When using a high-throughput ßuorescent microscope or plate-reader, assay for cell number in each well in order to meaningfully compare GFP signals across wells and plates. This can be done using a commercial cell viability assay, such as Resazurin Cell Viability Assay Kit (Biotium, cat. no. 30025) or the CellTiter-Glo Luminescent Cell Viability Assay (Promega, cat. no. G7570). The percentage of eGFP+ cells using this differentiation method should be similar to that obtained through EB differentiation (Basic Protocol 1). BASIC PROTOCOL 2 Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

PREPARATION OF DIFFERENTIATION CULTURE FOR FLOW CYTOMETRY ANALYSIS OR FACS This protocol describes the dissociation of EBs into a single-cell suspension to be used for ßow cytometry analysis or FACS. Dissociation is also recommended for isolating particular EBs or areas in the differentiation culture for long-term culture (see Basic Protocol 4).

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Materials EB culture (see Basic Protocol 1) 0.25% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25200-056) Collagenase/DNase solution (see recipe) HEPES-buffered saline (HBS; see recipe) Flow cytometry buffer (see recipe) Propidium iodide solution (1 mg/ml; Invitrogen, cat. no. P3566) 10-ml serological pipet or sterile cell scraper 15-ml or 50-ml conical tubes 1000-μl pipettor 37◦ C water bath 40-μm cell strainers Mechanically dissociate EBs 1. Using a 10-ml serological pipet, collect up the medium from an EB culture dish and eject it back onto the dish to dislodge EBs that have adhered to the bottom. Alternatively, gently dislodge attached EBs using a cell scraper. Be careful not to generate excess foam while pipetting. The EBs should detach after a few rounds of pipetting (Fig. 1F.10.2A). Of note, during the later stages of differentiation (e.g., beyond day 10), EBs become increasingly adherent to the plate and will require more vigorous pipetting to detach.

2. Using a 10-ml serological pipet, transfer detached EBs into a 15-ml or 50-ml conical tube and let stand for 3 to 4 min, until the EBs have settled to the bottom of the tube (Fig. 1F.10.2B). Carefully aspirate the medium above the EBs. Since the EBs are denser than the medium, it is usually not necessary to centrifuge the tubes.

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Figure 1F.10.2 Harvesting EBs for isolation and analysis of cardiac progenitor cells. (A) After dissociation from the culture plate, EBs (indicated by arrows) appear as large particles in suspension. The plate shown is at day 7 of differentiation. (B) The contents of the plate shown in (A) are transferred to a 15-ml conical tube, and the EBs are allowed to settle to the bottom. Larger tubes (e.g., 50-ml conical tube) may be used when pooling multiple plates. Embryonic and Extraembryonic Stem Cells

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Digest day 6 and younger EBs with trypsin 3. For each 15-cm dish equivalent of EBs, add 0.5 ml 0.25% trypsin/EDTA to the dislodged cells and pipet gently to mix. 4. Incubate tubes 5 min in a 37◦ C water bath. 5. Inactivate the trypsin with differentiation medium at a minimum of 4× the volume of enzyme used. Continue with step 9.

Digest day 7 and older EBs with collagenase/DNase 6. For each 15-cm dish equivalent of EBs, add 0.5 ml collagenase/DNase solution to the dislodged cells and pipet gently up and down 30 times with a 1000-μl pipettor to facilitate the breaking of EBs into a single-cell suspension. 7. Incubate tubes 1 hr in a 37◦ C water bath. Repeat trituration every 15 min during the incubation. 8. To remove collagenase/DNase solution after incubation, dilute with HBS at a minimum of 5 times the volume of the previously added collagenase/DNase solution.

Prepare the cells for ßow cytometry applications 9. Centrifuge the cells 3 min at 200 × g, room temperature. Aspirate the supernatant. 10. Resuspend the cell pellet in 0.5 ml ßow cytometry buffer for every 15-cm dish equivalent of cells, or in the appropriate medium or buffer for other applications. It may be necessary to dilute the cell suspension further to prevent clogging of the ßow cytometer or FACS machine. Adjust the volume of ßow cytometry buffer accordingly.

11. Add 1 μl of propidium iodide solution for every 0.5 ml buffer. Some visible clumps may remain after digestion and resuspension, especially during the later stages of differentiation. To remove these, Þlter the cell suspension through a 40-μm cell strainer.

BASIC PROTOCOL 3

FLOW CYTOMETRIC ANALYSIS AND FACS-BASED PURIFICATION OF CARDIAC PROGENITOR CELLS The Nkx2.5-eGFP marker allows for the quantiÞcation and puriÞcation of cardiac progenitor cells that develop during EB differentiation. Assessment of the efÞciency of cardiac differentiation can be performed using ßow cytometry to determine the percentage of cells that have committed to the cardiac lineage (based on the expression of eGFP). After cells have been prepared according to Basic Protocol 2, standard ßow cytometry procedures are used to quantify the percentage of eGFP+ cells with each batch of differentiation. A typical series of gates used for determining the percentage of eGFP+ cells present in differentiated EBs is shown in Figure 1F.10.3. We routinely observe (from day 7 of differentiation onward) 1% to 5% of eGFP+ cells present in the total PI-negative live-cell population. The exact yield varies from experiment to experiment, but within the same experiment, the percentage peaks at day 10 and remains more or less constant before diminishing at day 18, when the EB culture begins to degrade.

Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

Fluorescence-activated cell sorting (FACS) can be used to recover the eGFP+ cells for further culture and/or experimentation (see Basic Protocol 4), if desired.

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Figure 1F.10.3 Flow cytometry analysis of differentiated (day 10) EBs. A single-cell suspension of EB-derived differentiated Nkx2.5-eGFP cells were prepared according to Basic Protocol 2 and analyzed using a FACSCalibur ßow cytometer. (A) Forward- vs. side-scatter plot, gated to exclude noncellular debris. (B) Gating for live (PI-) cells. (C) Gating for GFP+ cells. The percentage of GFP+ cells in this particular analysis, 3.84%, is typical for a normal differentiation using healthy cells.

CULTURE AND FUNCTIONAL CHARACTERIZATION OF CARDIAC PROGENITORS

BASIC PROTOCOL 4

This protocol describes the continued culturing and characterization of the Nkx2.5eGFP+ cardiac progenitor cells isolated by FACS from EB differentiation. There are a wide variety of functional assays that may be used to characterize these progenitor cells and their progeny. We restrict ourselves here to describing how to prepare cells for gene expression analysis and immunocytochemistry; for more specialized techniques, such as electrophysiological assessment (Maltsev et al., 1994) or contractility assays (Pfannkuche et al., 2009), please consult the relevant literature.

Materials EBs (dissociated according to Basic Protocol 2) Differentiation medium (DM; see recipe) TRIzol (Invitrogen, cat. no. 15596026) or other cell lysis solution for RNA isolation

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Antibodies for cardiac troponin T (cTnT; e.g., clone 13-11, Neomarkers), sarcomeric myosin heavy chain (sarcMHC; e.g., clone MF20, Developmental Studies Hybridoma Bank), sarcomeric actinin (sarcActinin; e.g., clone EA-53, Sigma), smooth muscle actin-α (SMAα; e.g., clone IA4, DakoCytomation), and smooth muscle myosin heavy chain (SM-MHC; e.g., polyclonal, Biomedical Technologies) FACS collection tubes (Becton Dickinson, cat. nos. 352054, 352063, or 352196) Gelatin-coated chamber slides or 6-, 12-, 24-, or 96-well plates Culture Nkx2.5+ cardiac progenitors after isolation by FACS 1. During FACS isolation of Nkx2.5-eGFP+ cardiac progenitor cells, collect the eGFP+ cells directly into DM. For a single batch of twenty Þve 15-cm dishes, 3 ml of DM in a 5 ml FACS collection tube is sufÞcient to accommodate the 250,000 to 500,000 eGFP+ cells typically obtained.

2. After the collection is Þnished and provided that the volume of saline carried over from FACS is less than 10% of total medium volume, plate the cells directly into gelatin-coated chamber slides for immunocytochemistry, or into 6-, 12-, 24-, or 96well plates as required for further experimentation. Alternatively, if gene expression analysis is desired, pellet the cells by centrifugation for RNA preparations using TRIzol or similar reagents. We recommend a cell seeding density in the range of 40 to 400 cells/mm2 . If the volume of FACS saline carryover is >10%, centrifuge the cells and resuspend in fresh DM.

3. Depending on the experimental condition required and the density of cells in each well, feed the collected Nkx2.5+ cardiac progenitor cells every 2 to 3 days with fresh differentiation medium. If isolated at day 5 or 6 of EB differentiation, the eGFP+ cardiac progenitor cells will continue to differentiate spontaneously into cardiomyocytes (∼60%) or smooth muscle cells (∼40%).

Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

Figure 1F.10.4 Staining for the sarcomeric structures of ES-derived cardiomyocytes using antisarcomeric actinin mouse monoclonal antibody.

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On rare occasions, there may be CD31+ cells by gene expression or immunocytochemistry, but it is unclear if these represent true endocardial cells. The identity of differentiated progenies from puriÞed cardiac progenitor cells can be elucidated by antibody staining for early (sarc. MHC, SMAα) and late (cTnT, SM-MHC) markers for cardiomyocytes and smooth muscle cells, respectively. From differentiation cultures at day 10 and beyond, staining of cardiac progenitor cellderived cardiomyocytes with sarcomeric actinin will reveal the presence of a striation pattern characteristic for mature cardiomyocytes (Fig. 1F.10.4).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. Unless otherwise noted, all solutions and media are 0.2 μm-Þlter sterilized.

Collagenase/DNase solution Dissolve 50 mg each of collagenase A (Roche; cat. no. 11088785103) and collagenase B (Roche, cat. no. 11088823103) in 5 ml of ßow cytometry buffer (see recipe). Store up to 4 weeks at 4◦ C. Prior to use, supplement an aliquot of the required volume with DNase (Calbiochem, cat. no. 260913) to a Þnal concentration of 10 μg/ml.

Differentiation medium 500 ml IMDM (Invitrogen, cat. no.12440-061) 6.25 ml of 200 mM L-glutamine (Invitrogen, cat. no. 25030-081; Þnal concentration 2 mM) 94 ml fetal bovine serum (FBS; Invitrogen, cat. no. 10437-028; Þnal concentration 15% v/v) 6.5 μl monothioglycerol (Sigma, cat. no. M6145) 6.25 ml 5 mg/ml ascorbic acid (aqueous solution; Sigma, cat. no. A4544; Þnal concentration 50 μg/ml) Store up to 2 weeks at 4◦ C L-glutamine is very sensitive to oxidation and must be replenished every 7 days. FBS should be lot-tested for optimal cardiac differentiation results.

DMEM-ES medium 500 ml DMEM-high glucose (Invitrogen, cat. no. 11965-0692) 6.25 ml of 100 mM non-essential amino acids (Invitrogen, cat. no. 11140-050; Þnal concentration 1 mM) 6.25 ml of 200 mM L-glutamine (Invitrogen, cat. no. 25030-081) 12.5 ml penicillin/streptomycin (10,000 U/ml, 10,000 μg/ml; Invitrogen, cat. no. 15070-063; Þnal concentration 200 U/ml penicillin, 200 μg/ml streptomycin) 94 ml fetal bovine serum (FBS; GEMINI, cat. no. 100-106) 4.4 μl 2-mercaptoethanol (Sigma, cat. no. M6250) 62.5 μl leukemia inhibitory factor (107 units/ml; CHEMICON, cat. no. ESG1107; Þnal concentration 10,000 U/ml) Store up to 2 weeks at 4◦ C L-glutamine

is very sensitive to oxidation and must be replenished every 7 days.

FBS should be lot-tested for minimal induction of differentiation.

Flow cytometry buffer Add fetal bovine serum (GEMINI; cat. no. 100-106) to HBS (see recipe) solution to a Þnal FBS concentration of 20%. Store up to 3 months at 4◦ C.

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Gelatin, 0.1% (w/v) Dissolve 0.5 g of gelatin (from porcine skin) in 500 ml distilled/deionized water and autoclave. Store indeÞnitely at room temperature.

Gelatinized plates Dispense 0.1% gelatin solution (see recipe) into tissue culture vessels to cover the growth surface and incubate 15 min at room temperature. At the end of incubation, aspirate excess gelatin solution and let the tissue culture plate surface dry for 5 min. Store up to 1 to 2 months at 4◦ C until use.

HEPES-buffered saline (HBS) 2.5 ml of 1 M HEPES (Invitrogen, cat. no. 15630-080) 2.68 ml of 5 M NaCl 0.5 ml of 1 M KCl 70 μl of 1 M Na2 HPO4 94.25 ml ddH2 O Adjust pH to 7.1 using 1 N NaOH Store up to 6 months at 4◦ C IMDM-ES medium 500 ml IMDM (Invitrogen, cat. no.12440-061) 6.25 ml of 200 mM L-glutamine (Invitrogen, cat. no. 25030-081) 12.5 ml penicillin/streptomycin (10,000 U/ml, 10,000 μg/ml; Invitrogen, cat. no. 15070-063) 94 ml fetal bovine serum (FBS; GEMINI, cat. no. 100-106) 6.5 μl monothioglycerol (Sigma, cat. no. M6145) 62.5 μl leukemia inhibitory factor (107 units/ml; CHEMICON, cat. no. ESG1107) Store up to 2 weeks at 4◦ C L-glutamine

is very sensitive to oxidation and must be replenished every 7 days.

FBS should be lot-tested for minimal induction of differentiation.

COMMENTARY Background Information

Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

The self-renewal and developmental potential of pluripotent stem cells make them an attractive source for rare cell populations that arise during embryonic development, such as the earliest organ-speciÞc progenitor populations, from which all or a large subset of cell types of a particular organ are derived. It has been demonstrated that such a population exists for the mammalian heart and is marked by positive Nkx2.5, Isl1, and/or Flk-1 expression in mice (Kattman et al., 2006; Moretti et al., 2006; Wu et al., 2006) and KDR-1 or Isl1 expression in differentiated human ES cells (Yang et al., 2008; Bu et al., 2009). These progenitor cells give rise to cardiomyocytes, smooth muscle cells (Wu et al., 2006; Bu et al., 2009), and endothelial cells (Bu et al., 2009), the three primary cell types of the heart. Although cellular markers for isolation of cardiac progenitor cells from in vivo and in vitro

contexts have now been identiÞed, the process by which a pluripotent stem cell Þrst becomes committed to a multipotent lineage progenitor remains largely unknown. Further inquiry into these processes holds vast potential for clinical translation, particularly in the areas of congenital heart disease and regenerative therapy for myocardial injury.

Critical Parameters The developmental potential of ES or iPS cells is strongly affected by a number of different factors, the most important of which is their ability to remain uniformly undifferentiated prior to initiating differentiation. During propagation, the cells should always be maintained in fresh medium and be passaged before reaching 80% conßuency in the culture dish. As the passage number increases (>30), the differentiation capacity of ES and iPS cells declines (Nagy et al., 1990, 1993; Wang et al., 1997). It

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is advisable to use lower-passage cells for experimentation. Cells that remain undifferentiated should exhibit small size and round shape with large nuclear-to-cytoplasmic ratio. When cultured on feeder MEFs, they should aggregate into larger colonies with raised borders and refractile surfaces. They should be free of any contamination, including mycoplasma. Typical signs of poor quality cells include the appearance of large numbers of cellular fragments ßoating in the culture medium, the presence of vacuoles on cell surface or within each cell, and colonies with irregular borders and ßattened cell morphology. If the culture medium turns yellow or yellow-orange in color earlier than usual, a suspicion should be raised for the presence of mycoplasma contamination. It has been our experience that cells exhibiting these defects do not differentiate well and should be discarded. The efÞciency of differentiation into the cardiac lineage is also inßuenced by the particular lot of fetal bovine serum used to make differentiation medium. Several different lots should be tested, and the one that yields the highest percentage of eGFP+ cells at days 7 to 10 of differentiation should be used exclusively for future experiments. Finally, cell density at the outset of differentiation is also an important factor contributing to the yield of cardiac progenitors. For both EB and monolayer differentiation protocols, the stated number of cells per droplet or well, respectively, have been empirically determined to maximize the number of eGFP+ cells obtained. In adapting the protocols of this unit for ES or iPS lines carrying different transgenic cardiac markers, it may be necessary to adjust the cell seeding density for optimal differentiation.

Anticipated Results The protocols outlined in this unit can be scaled up to yield large numbers (105 –106 ) of Nkx2.5+ cardiac progenitors for a variety of applications directed towards investigating early cardiac development and the therapeutic potential of progenitor populations and their derivatives. Upon continued culture, Nkx2.5+ progenitor cells should spontaneously differentiate into beating cardiomyocytes and smooth muscle cells.

of differentiation. We Þnd that past day 18, formerly beating clusters tend to stop beating, and eGFP expression decreases. A typical high-throughput screen experiment takes up to 9 days, 2 days again for MEF depletion, followed by 1 week of monolayer differentiation. Beyond 1 week, the cells become too dense to dissociate easily for ßow cytometry, and their further development is inhibited by nutrient limitation. Preparation of a full batch of twenty Þve 15-cm EB plates for FACS requires 3 to 4 hr, followed by an additional 2 to 4 hr on the FACS machine.

Literature Cited Bu, L., Jiang, X., Martin-Puig, S., Caron, L., Zhu, S., Shao, Y., Roberts, D.J., Huang, P.L., Domian, I.J., and Chien, K.R. 2009. Human Isl1 heart progenitors generate diverse multipotent cardiovascular cell lineages. Nature 460:113-117. Domian, I.J., Chiravuri, M., van der Meer, P., Feinberg, A.W., Shi, X., Ying, C., Wu, S., Parker, K., and Chien, K.R. 2009. Committed ventricular progenitors in the Islet-1 lineage expand and assemble into functional ventricular heart muscle. Science 326:426-429. Kattman, S.J., Huber, T.L., and Keller, G.M. 2006. Multipotent ßk-1+ cardiovascular progenitor cells give rise to the cardiomyocyte, endothelial, and vascular smooth muscle lineages. Dev. Cell 11:723-732. Lints, T.J., Parsons, L.M., Hartley, L., Lyons, I., and Harvey, R.P. 1993. Nkx-2.5: A novel murine homeobox gene expressed in early heart progenitor cells and their myogenic descendents. Development 119: 419-431. Lyons, I., Parsons, L.M., Hartley, L., Li, R., Andrews, J.E., Robb, L., and Harvey, R.P., 1995. Myogenic and morphogenetic defects in the heart tubes of murine embryos lacking the homeo box gene Nkx2-5. Genes Dev. 9:16541666. Maltsev, V.A., Wobus, A.M., Rohwedel, J., Bader, M., and Hescheler, J. 1994. Cardiomyocytes differentiated in vitro from embryonic stem cells developmentally express cardiac-speciÞc genes and ionic currents. Circ. Res. 75:233-244. Moretti, A., Caron, L., Nakano, A., Lam, J.T., Bernshausen, A., Chen, Y., Qyang, Y., Bu, L., Sasaki, M., Martin-Puig, S., Sun, Y., Evans, S.M., Laugwitz, K.L., and Chien, K.R. 2006. Multipotent embryonic isl1+ progenitor cells lead to cardiac, smooth muscle, and endothelial cell diversiÞcation. Cell 127:1151-1165.

Time Considerations

Nagy, A., G´ocza, E., Merentes Dias, E., Prideaux, V.R., Iv´anyi, M., and Rossant, J. 1990. Embryonic stem cells alone are able to support fetal development in the mouse. Development 110:815821.

A typical EB differentiation experiment requires 9 to 20 days, beginning with 2 days for MEF depletion, followed by 1 to 2 weeks

Nagy, A., Rossant, J., Nagy, R., AbremowNewerly, W., and Roder, J.C. 1993. Derivation of completely cell culture-derived mice from

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early-passage embryonic stem cells. Proc. Nat. Acad. Sci. U.S.A. 90:8424-8428. Pfannkuche, K., Liang, H., Hannes, T., Xi, J., Fatima, A., Nguemo, F., Matzkies, M., Wernig, M., Jaenisch, R., Pillekamp, F., Halbach, M., Schunkert, H., Saric, T., Hescheler, J., and Reppel, M. 2009. Cardiac myocytes derived from murine reprogrammed Þbroblasts: Intact hormonal regulation, cardiac ion channel expression and development of contractility. Cell Physiol. Biochem. 24:73-86. Samuelson, L. and J.M. Metzger. 2006. Differentiation of embryonic stem (ES) cells using the hanging drop method. Cold Spring Harbor Protocols doi:10.1101/pdb.prot4485. Stanley, E.G., Biben, C., Elefanty, A., Barnett, L., Koentgen, F., Robb, L., and Harvey, R.P. 2002. EfÞcient Cre-mediated deletion in cardiac progenitor cells conferred by a 3 UTR-ires-Cre allele of the homeobox gene Nkx2-5. Int. J. Dev. Biol. 46:431-439. Takahashi, T., Lord, B., Schulze, P.C., Fryer, R.M., Sarang, S., Gullans, S., and Lee, R.T. 2003. Ascorbic acid enhances differentiation of embryonic stem cells into cardiac myocytes. Circulation 107:1912-1916.

Tremml, G., Singer, M., and Malavarca, R. 2008. Culture of mouse embryonic stem cells. Curr. Protoc. Stem Cell Biol. 5:1C.4.1-1C.4.19. Wang, Z., Kiefer, F., Urb´anek, P., and Wagner, E.F. 1997. Generation of completely embryonic stem cell-derived mutant mice using tetraploid blastocyst injection. Mech. Dev. 62:137-145. Wobus, A.M., Wallukat, G., and Hescheler, J. 1991. Pluripotent mouse embryonic stem cells are able to differentiate into cardiomyocytes expressing chronotropic responses to adrenergic and cholinergic agents and Ca2+ channel blockers. Differentiation 48:173-182. Wu, S.M., Fujiwara, Y., Cibulsky, S.M., Clapham, D.E., Lien, C., Schultheiss, T., and Orkin, S.H. 2006. Developmental origin of a bipotential myocardial and smooth muscle cell precursor in the mammalian heart. Cell 127:11371150. Yang, L., Soonpaa, M.H., Adler, E.D., Roepke, T.K., Kattman, S.J., Kennedy, M., Henckaerts, E., Bonham, K., Abbott, G.W., Linden, R.M., Field, L.J., and Keller, G.M. 2008. Human cardiovascular progenitor cells develop from a KDR+ embryonic stem cell-derived population. Nature 453:524-528.

Isolation and Functional Characterization of Pluripotent Stem Cell-Derived Cardiac Progenitor Cells

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Differentiation of Mouse Embryonic Stem Cells into Cardiomyocytes via the Hanging-Drop and Mass Culture Methods

UNIT 1F.11

Christopher J. Fuegemann,1 Ajoy K. Samraj,2 Stuart Walsh,2 Bernd K. Fleischmann,1 Stefan Jovinge,2,3 and Martin Breitbach1 1

Institute of Physiology I, Life & Brain Center, University of Bonn, Bonn, Germany Lund Strategic Research Center for Stem Cell Biology and Cell Therapy, Lund, Sweden 3 Department of Cardiology, Lund University Hospital, Lund Strategic Research Center for Stem Cell Biology and Cell Therapy, Lund, Sweden 2

ABSTRACT Herein, we describe two protocols for the in vitro differentiation of mouse embryonic stem cells (mESCs) into cardiomyocytes. mESCs are pluripotent and can be differentiated into cells of all three germ layers, including cardiomyocytes. The methods described here facilitate the differentiation of mESCs into the different cardiac subtypes (atrial-, ventricular-, nodal-like cells). The duration of cell culture determines whether preferentially early– or late–developmental stage cardiomyocytes can be obtained preferentially. This approach allows the investigation of cardiomyocyte development and differentiation in vitro, and also allows for the enrichment and isolation of physiologically intact cardiomyocytes for transplantation purposes. Curr. Protoc. Stem Cell Biol. 15:1F.11.1C 2010 by John Wiley & Sons, Inc. 1F.11.13.  Keywords: mouse embryonic stem cells (mESCs) r differentiation r hanging drops r mass culture r embryoid bodies r cardiomyocytes

INTRODUCTION In this unit, we describe two straightforward alternative protocols for the in vitro differentiation of mESCs into cardiomyocytes: the hanging-drop method (Basic Protocol 1), characterized by high reproducibility, and the mass culture method (Basic Protocol 2), better suited for harvesting large numbers of cardiomyocytes. Protocols for freezing (Support Protocol 1), thawing (Support Protocol 2), and culturing (Support Protocol 3) mESCs prior to their in vitro differentiation are included, all of which are critical steps for optimal performance of the cells during the differentiation period. Additionally, a protocol for feeder-cell depletion is supplied (Support Protocol 4). Feeder cells (mouse embryonic fibroblasts, MEFs) provide a matrix and nutrients for mESC growth in culture, but are known to influence the in vitro differentiation characteristics of pluripotent stem cells. NOTE: All procedures are performed in a class II biological hazard flow hood or a laminar-flow hood. NOTE: Solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator.

IN VITRO DIFFERENTIATION OF mESCs VIA HANGING DROPS This protocol is used for the differentiation of mESCs and is based on the method originally published by Wobus et al. (1991), with several modifications. In vitro differentiation into cardiomyocytes using this protocol, in our experience, results in better Current Protocols in Stem Cell Biology 1F.11.1-1F.11.13 Published online December 2010 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc01f11s15 C 2010 John Wiley & Sons, Inc. Copyright 

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Figure 1F.11.1 Differentiation of mESC in EBs. (A) Using the hanging-drop protocol, EBs at day 3 of differentiation are small and relatively homogenous in size. (B) EBs at day 10 of differentiation are larger and become more compact, but their growth is slowed down. (C) Using the mass culture protocol, EBs at day 10 of differentiation are less homogenous in size, shape, and structure. Scale bar = 200 μm.

reproducibility than that obtained with the mass culture protocol (Basic Protocol 2). However, the hanging-drop protocol is more time consuming and cost intensive in relation to total cardiomyocyte yield. Embryoid bodies (EBs) are generated by a defined number of cells, resulting in homogenous size and differentiation characteristics (Fig. 1F.11.1). Therefore, this method should preferentially be used for experiments requiring low biological variability, e.g., comparing differentiation under the influence of certain chemicals or conditions. Differentiation in hanging drops is achieved by first trypsinizing mESCs (see Support Protocol 3) and optionally depleting the feeder cell contamination to obtain an enriched mESC population (see Support Protocol 4). Next, mESCs are spotted in droplets onto the lid of an ultra-low-attachment dish and incubated upside down for 2 days. After 2 days, cell aggregates have formed, the so-called EBs (Fig. 1F.11.1). Figure 1F.11.2 shows the order of actions for the hanging-drop protocol schematically.

Materials PBS without Mg2+ or Ca2+ (CMF-PBS; Invitrogen, cat. no. 18912-014), sterile Pluripotent mESCs (Support Protocol 3) mESC differentiation medium (see recipe) Bacterial dishes or ultra-low-attachment dishes (Greiner, cat. no. 633180), 100-mm diameter Omni Trays (optional; Nunc, cat. no. 242811) Horizontal shaker 100-mm diameter tissue culture dishes (BD Falcon, cat. no. 35-3003), gelatin-coated (see recipe) or noncoated Additional reagents and equipment for culturing mESCs, including trypsinization (Support Protocol 3), for feeder cell depletion (Support Protocol 4), and for determination of viable cell number (UNIT 1C.3) Prepare dishes and cells 1. Prepare a sufficient number of 100-mm ultra-low-attachment dishes or Omni Trays with CMF-PBS (10 ml for the dishes or 20 ml for the Omni Trays) to ensure humidity and prevent the hanging drops from drying out. Differentiation of Mouse ESCs into Cardiomyocytes

Omni Trays resemble multiwell plates without wells, enabling the use of a multichannel pipettor to spot several droplets in parallel; their surface is about 1.5 times as large as a 100-mm dish. Per dish, approximately 100 EBs are generated, per Omni Tray 130 EBs.

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Hanging Drops

Mass Culture

D0 drops of ESC suspension on the lid of a bacterial dish cultured upside down

D0 cell suspension of defined ESC concentration cultured on a shaker

D2 EBs formed in hanging drops EBs are transferred into suspension culture

D2 EBs formed in suspension amount of EBs per ml is reduced by dilution

D7 EBs have grown in culture plating possible, or culturing is continued in suspension culture

D5 EBs have grown in culture plating possible, or culturing is continued in suspension culture

Figure 1F.11.2 Scheme of differentiation protocols. Flow chart showing schematically the order of actions for the hanging-drop (left) and mass culture (right) protocol.

2. Trypsinize cultured mESCs (see Support Protocol 3) and resuspend cells in 1 to 3 ml mESC differentiation medium. One 25-cm2 flask yields about 2–3 × 106 mESCs, which is sufficient to generate several thousand EBs.

3. Optional: Perform feeder cell depletion (see Support Protocol 4). 4. Determine viable cell number (UNIT 1C.3). If no feeder cell depletion has been performed, the number of feeder cells in the suspension is subtracted from the total cell number. To estimate the number of feeder cells, trypsinized cells of a similar-sized culture flask containing only feeder cells are counted. The feeder cell number is ∼60% of the seeding number, i.e., ∼7 × 105 for a 25-cm2 culture flask.

Plate cells 5. Dilute the suspension to 20,000 cells per ml with mESC differentiation medium and mix regularly in order to prevent the cells from sedimenting or clumping. 6. Pipet 20 μl-droplets each containing ∼400 mESC onto the lid of an ultra-lowattachment dish (up to 100 droplets can be accommodated) or on the lid of an

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Omni Tray (up to 130 droplets can be accommodated). Carefully turn the lid with the droplets upside down on top of a dish or tray previously filled with CMF-PBS (step 1). Incubate 48 hr. This day is defined as day 0 of differentiation (D0).

7. After 48 hr of incubation (differentiation day 2), collect the formed EBs by flushing the droplets with 5 ml of fresh mESC differentiation medium. Alternatively, the lid of the dish can be knocked carefully on the surface of a bench to collect the droplets at the rim of the dish.

8. Transfer the EBs into a new bacterial or ultra-low-attachment dish with the medium used for flushing the droplets, and fill up to 10 ml per dish. EBs from two 100-mm dishes are pooled into one dish, and EBs of three Omni Trays are distributed into two dishes, resulting in up to 200 EBs per dish.

9. Incubate EBs on a horizontal shaker at a frequency of 60 to 70 rpm to prevent clumping.

Feed cells 10. Change two-thirds of the medium every second day by carefully tilting the dish so that the EBs sink to the bottom before removing the old medium. Alternatively, one can swirl the dishes slowly to gather EBs in the middle of the dish and aspirate the medium from the side of the dish. Swirling is the best way to gather EBs at one spot for documentation and/or analysis purposes.

11. Optional: At day 7 of differentiation, plate EBs on gelatin-coated or noncoated 100-mm tissue culture dishes, which may enhance differentiation toward the cardiac lineage. EBs can also be kept in suspension but, in our experience, plating enhances differentiation toward the cardiac lineage; the underlying mechanism is not understood. Attachment of EBs makes medium changes more convenient. However, one has to keep an eye on floating EBs that have not attached to the surface and that might be aspirated with the old medium.

Monitor cultures for differentiation 12. At day 8 to 9 of differentiation, begin monitoring EBs for beating areas. A straightforward way of evaluating cardiogenic differentiation is to count EBs with beating areas. Normally, spontaneous beating is visible as early as day 7 of differentiation in 40% to 100% of the EBs, depending on the specific cell lines/clones. Each beating area scores for cardiogenic differentiation in an EB; note that this area can be relatively small compared to total EB size. Cardiogenic differentiation can be characterized in more detail using PCR and/or immunocytochemical staining. For adequate cardiac markers, see Anticipated Results. EBs can be maintained for several weeks in culture with medium changes every 2 to 3 days. However, note that cardiomyocyte yield does not further increase after day 12, due to the slowdown of proliferation (see Anticipated Results). Cardiomyocytes can be enriched by cutting out beating areas with a scalpel. This should be done under sterile conditions using a microscope under the flow hood. Cells can then be separated by gentle trypsinization (according to Support Protocol 2). A more convenient approach for purification of cardiomyocytes is the use of genetically manipulated mESCs, as described in the Background Information. Differentiation of Mouse ESCs into Cardiomyocytes

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IN VITRO DIFFERENTIATION OF mESCs VIA MASS CULTURE The method described in detail here is a culture protocol originally published by Kolossov et al. (2006), with several modifications. In vitro differentiation of mESCs with this method yields a higher number of cardiomyocytes and is less labor-intensive than the hanging-drop method. However, reproducibility is not as good as with the hanging-drop method because there is more variation in EB size (Fig. 1F.11.1C) and in the yield of cardiomyocytes at different days of differentiation. Thus, mass culture is preferable if relatively large amounts of cardiomyocytes are required, i.e., for protein-expression studies or transplantation purposes.

BASIC PROTOCOL 2

For differentiation in mass culture, mESCs are firstly trypsinized (see Support Protocol 3) and optionally depleted from feeder cell contamination to get a relatively pure mESC population (see Support Protocol 4). Then, a single-cell suspension with a defined number of mESCs in differentiation medium is incubated for 2 days on a shaker to allow formation of EBs. Figure 1F.11.2 shows the order of actions for the mass culture protocol schematically.

Materials Pluripotent mESCs (Support Protocol 3) PBS without Mg2+ or Ca2+ (CMF-PBS; Invitrogen, cat. no. 18912-014), sterile mESC differentiation medium (recipe see below) Bacterial dishes or ultra-low-attachment dishes (Greiner, cat. no. 633180), 100-mm diameter Omni Trays (optional; Nunc, cat. no. 242811) Horizontal shaker: Gesellschaft f¨ur Labortechnik mbH (GFL, http://www.gfl.de/) 50-ml conical tubes Microscope: 5×, 10×, and 20× oculars 100-mm diameter tissue culture dishes (BD Falcon, cat. no. 35-3003), gelatin-coated (see recipe) or noncoated Additional reagents and equipment for culturing mESCs, including trypsinization (Support Protocol 3), for feeder cell depletion (Support Protocol 4), and for determination of viable cell number (UNIT 1C.3) Prepare cells 1. Trypsinize cultured mESCs (see Support Protocol 3) and resuspend in 1 to 3 ml mESC differentiation medium. 2. Optional: Perform feeder cell depletion (see Support Protocol 4). 3. Determine viable cell number (UNIT 1C.3). If no feeder cell depletion has been performed, the number of feeder cells in the suspension is subtracted from the total cell number. One 25-cm2 flask yields about 2–3 × 106 mESCs, which is sufficient to generate two to three mass culture dishes. To estimate the number of feeder cells, trypsinized cells of a similar-sized culture flask containing only feeder cells are counted. The feeder cell number is ∼60% of the seeding number, i.e., ∼7 × 105 for a 25-cm2 culture flask.

Generate EBs 4. To generate EBs, suspend 1 × 106 mESCs in 10 ml of mESC differentiation medium and incubate the suspension in a 100-mm bacterial or ultra-low-attachment dish on a horizontal shaker at a shaking velocity of 60 to 70 rpm. This day is defined as differentiation day 0 (D0).

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5. After 48 hr of incubation (differentiation day 2), determine the number of EBs in the suspension. 6. For counting, collect EBs in medium in a 50-ml conical tube. Invert tube frequently. EBs are quite heavy aggregates and sink to the bottom of the tube quickly. Therefore, the suspension is carefully inverted frequently for equal distribution of EBs.

7. Pipet 20 μl of EB suspension onto a lid of a dish for counting, and count using 500× to 1000× magnification. For easy counting, the EB suspension is drawn as a little line of fluid instead of spotting as a droplet. Counting two or three samples is recommended.

Maintain EBs 8. Prepare appropriate numbers of 100-mm bacterial or ultra-low-attachment dishes with 10 ml of mESC differentiation medium. 9. After counting, transfer 500 to 1000 EBs into each dish and incubate further on a horizontal shaker at a frequency of 60 to 70 rpm. Under these conditions differentiating EBs grow in suspension and will not attach.

10. Change two-thirds of the medium every second day, as described in Basic Protocol 1. 11. Optional: At day 5 of differentiation, plate EBs on gelatin-coated or noncoated 100mm diameter tissue culture dishes, which may enhance differentiation toward the cardiac lineage. In the authors’ hands, plating at day 5 rather than at day 7 of differentiation results in more pronounced cardiogenic differentiation when using the mass culture protocol.

Monitor EBs for differentiation 12. At day 8 to 9 of differentiation, begin monitoring EBs for beating foci (see Basic Protocol 1). EBs can be maintained for several weeks in culture with medium changes every 2 to 3 days. However, note that cardiomyocyte yield does not further increase after day 12, due to the slowdown of proliferation (see Anticipated Results). Cardiomyocytes can be enriched by cutting out beating areas with a scalpel. This should be done under sterile conditions using a microscope under the flow hood. Cells can then be separated by gentle trypsinization (according to Support Protocol 2). A more convenient approach for purification of cardiomyocytes is the use of genetically manipulated mESCs, as described in the Background Information. SUPPORT PROTOCOL 1

FREEZING mESCs mESCs are preserved well in liquid nitrogen as suspensions containing 50% fetal bovine serum and 10% DMSO. The cell concentration is not crucial for viability of cells; it usually ranges between 0.3–5 × 106 cells/ml.

Materials Cultures of mESC mESC culture medium (see recipe) Freezing medium: 80% (v/v) FBS/20% (v/v) DMSO, ice cold Liquid N2 Differentiation of Mouse ESCs into Cardiomyocytes

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Cryotubes (Nunc, cat. no. 375418) 15- and 50-ml conical tubes Freezing container (Nalgene 5100-0001) Liquid nitrogen container Current Protocols in Stem Cell Biology

Additional reagents and equipment for trypsinizing mESCs (Support Protocol 2) and determining viable cell number (UNIT 1C.3) 1. Trypsinize cultured mESCs (see Support Protocol 2), and resuspend cells in 1 ml of mESC culture medium. 2. Determine viable cell number (UNIT 1C.3), as described above. 3. Label the cryotubes intended for use. 4. Add mESC culture medium to achieve a cell concentration of 2 × 106 cells/ml (to freeze 1 × 106 cells per cryotube). The cell concentration should be twice as high as intended for freezing the cells. The cell number to transfer into a cryotube depends on the experiments planned and on the specific clone. Fast-growing clones can be kept in smaller batches than slow-growing ones. Additionally, for long-term storage, freeze the cells at a higher concentration, because some cells will die over time, even when stored in liquid nitrogen.

5. Add an equal volume of ice-cold freezing medium to the cell suspension, gently mix, and transfer the diluted suspension into the prepared cryotubes, 1 ml per tube. If a lot of cryotubes need to be frozen, it is beneficial for cell survival to put the already filled cryotubes directly on ice.

6. Quickly transfer the cryotubes into the freezing container and put it into a –80◦ C freezer. The freezing device will slowly cool the probe down to prevent the formation of sharp ice crystals.

7. On the next day, transfer the cryotubes into the liquid nitrogen container.

THAWING mESCs Cells are quickly thawed in order to prevent damage caused by prolonged DMSO exposure. Defined numbers of mESCs are then seeded onto mitotically inactivated feeder cells or gelatin-coated culture dishes, which allow mESCs to attach and expand (Support Protocol 3). Before thawing cryotubes containing mESCs, culturing medium is warmed up in a 37◦ C water bath to speed up thawing of the frozen cells.

SUPPORT PROTOCOL 2

Materials Pluripotent mouse embryonic stem cells (mESCs; Support Protocol 3), frozen (Support Protocol 1) mESC culture medium (see recipe) MEFs, mitotically inactivated (γ irradiated or mitomycin C treated; UNIT 1C.3) 15- and 50-ml conical centrifuge tubes Centrifuge 25-cm2 tissue culture flasks Additional reagents and equipment for counting viable cells (UNIT 1C.3) 1. Transfer cryotubes containing frozen mESCs into a 37◦ C water bath straight from the liquid nitrogen tank. 2. Immediately after thawing, transfer the cell suspension into a 15-ml conical tube containing prewarmed mESC culture medium. The volume of culture medium should be at least three times the volume of freezing solution, to minimize the negative effect of DMSO on cell viability.

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3. Centrifuge the tube containing cell suspension for 5 min at 250 × g, room temperature, to collect the cells. 4. Discard supernatant and resuspend cell pellet in 1 ml of fresh, prewarmed mESC culture medium. 5. Count viable cells (UNIT 1C.3). In the authors’ experience, the percentage of dead cells increases with storage time, especially when cells are stored under suboptimal conditions, e.g., in the freezer at −80◦ C.

6. Seed the cells in 25-cm2 flasks at 1–3 × 105 cells/flask. The number of cells used for seeding depends upon the cell line being used.

7. Incubate. SUPPORT PROTOCOL 3

CULTURING mESCs Herein, we describe the culture conditions necessary for the long-term propagation of mESCs. This procedure is aimed at preventing differentiation and reducing chromosomal aberrations of mESCs, which otherwise may occur within several passages (Nagy et al., 1993). Prior to starting, appropriate numbers of flasks or dishes containing a layer of mitotically inactivated feeder cells are established. The inactivated feeder cells are seeded into the tissue culture flask at least 24 hr prior to plating of mESCs, to facilitate their attachment and allow their expansion to occupy as much of the surface area of the flask as possible. Feeder cells are prepared at a concentration of 5 × 105 /ml and seeded at 1 ml per 10 cm2 (use 2.5 ml for a 25-cm2 and 7.5 ml for a 75-cm2 flask, respectively).

Materials Cultures of mESC in 25-cm2 or 75-cm2 flasks Phosphate-buffered saline, Ca2+ and Mg2+ free (CMF-PBS; Invitrogen, cat. no. 18912-014), sterile Trypsin-EDTA (Invitrogen, cat. no. 25300) mESC culture medium (see recipe) Mitotically inactivated MEF feeder cells (UNIT 1C.3) 15- and 50- ml conical tubes (e.g., BD Falcon) Centrifuge 25-cm2 tissue culture flasks Additional reagents and equipment for preparing mitotically inactivated MEF feeder layers (UNIT 1C.3) 1. Wash the cultured mESC colonies once with 10 ml CMF-PBS. 2. Add enough trypsin/EDTA to cover the entire surface (1 ml/3 ml of trypsin for 25-cm2 /75-cm2 flasks, respectively) and incubate at 37◦ C for 3 to 5 min until cells detach. Detachment of cells can be periodically monitored under the microscope to prevent over-digestion and cell damage.

3. Stop the trypsinization reaction by adding at least a 3-volume excess of mESC culture medium. Differentiation of Mouse ESCs into Cardiomyocytes

4. Gently pipet up and down to obtain a single-cell suspension.

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Figure 1F.11.3 Culturing of mESC colonies. (A) Early, small colonies are characterized by a roundish shape with sharp borders in a phase-contrast microscope. (B) Grown colonies have to be passaged before the rim begins to assume an irregular shape or colonies start to become confluent. (C) When not passaged in time, mESC colonies become flat, spread out, and appear integrated within the feeder layer. Over time the typical shape and contrast of the colonies is entirely lost. Scale bar = 50 μm.

5. Transfer this to a 15- or 50-ml conical tube and centrifuge for 5 min at 250 × g, room temperature. 6. Discard supernatant and resuspend pellet in 1 to 3 ml fresh, prewarmed mESC culture medium. 7. Transfer cell solution into a flask pre-seeded (UNIT 1C.3) with feeder cells, e.g., 3 × 105 mESCs in 4 ml of mESC culture medium in a 25-cm2 flask. However, the number of mESCs to seed a single flask depends on the growth rate of the respective clone and needs to be established for each individual cell line/clone. Normally, it ranges between 1 × 105 and 3 × 105 mESCs in a 25-cm2 flask. In order to determine the exact mESC number, the number of feeder cells is subtracted from the total cell count. If applicable, mESC clones can be maintained without the need for a feeder layer and can be seeded on gelatin-coated cell culture dishes (many mESC lines grow on gelatin, e.g., D3, G4, and R1). When mESCs are grown on gelatin-coated surfaces, twice the normal amount of LIF (2000 U/ml) is added to the culture medium to compensate for the lack of feeder-provided LIF in the culture.

8. Make sure that the cells are spread evenly by carefully moving the flask on an even surface in circles mimicking the number “8”. 9. In order to prevent the cells from differentiating, monitor colony growth very carefully by using a phase-contrast microscope at 500× to 1000× magnification. 10. Split the mESCs regularly before colonies become confluent and start to lose their distinct roundish shape (Fig. 1F.11.3). If colonies have not grown large enough to split 2 days after passaging, replace culture medium every second day until colonies have grown large enough (colonies for passaging should be at least as large as shown in Fig. 1F.11.3A, i.e., 50-μm diameter).

DEPLETION OF FEEDER CELLS (“PRE-PLATING”) The presence of feeder cells is known to affect the in vitro differentiation of mESCs in EBs. The pre-plating procedure described here is a simple and efficient approach to reduce the number of contaminating feeder cells. Before starting the trypsinization, the required number of 100-mm tissue culture dishes are coated with 5 ml of 0.1% gelatin as described in the Reagents and Solutions. One dish is sufficient for two 25-cm2 culture flasks.

SUPPORT PROTOCOL 4

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Materials Culture of mESCs mESC culture medium (see recipe) 100-mm gelatin-coated tissue culture dishes (see recipe) 15-ml conical centrifuge tubes Additional reagents and equipment for culturing mESCs, including trypsinization (Support Protocol 3) 1. Trypsinize cultured mESC (Support Protocol 3). 2. Resuspend the pellet in fresh mESC culture medium at a final concentration of 1–3 × 106 cells/ml. 3. Plate up to 5 × 106 cells in 10 ml mESC culture medium onto a gelatin-coated 100-mm dish and incubate for 20 min at 37◦ C. During this time, the majority of feeder cells will attach while mESCs will mainly remain in suspension.

4. Collect mESCs with a pipet by gently tilting the dish, and transfer cell suspension to a 15-ml conical tube. 5. For differentiation experiments, directly proceed with the Basic Protocols. Alternatively, the cells are cultivated on gelatin-coated dishes for 2 days followed by another pre-plating step to further reduce the number of remaining feeder cells. Note that this is only recommended for mESC lines that can be maintained on gelatin.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Gelatin solution 0.1% (w/v) Dissolve 0.5 g of gelatin powder in 0.5 liter CMF-PBS (Invitrogen, cat. no. 18912014) and autoclave. Solution can be stored for several months at room temperature.

Gelatin-coated flasks/dishes For cultivation of mESCs and feeder depletion, tissue culture dishes or flasks are coated with a sufficient volume of 0.1% gelatin solution (see recipe) to completely cover the entire surface of the dishes or flasks. The gelatin-coated dishes/flasks can be stored at 4◦ C for several weeks when sealed with parafilm to prevent their drying out. After incubation for a minimum of 30 min at 37◦ C, gelatin is aspirated and excess gelatin on the dishes/flasks removed by washing with CMF-PBS (Invitrogen, cat. no. 18912-014) or medium before seeding cells.

mESC culture medium

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415 ml DMEM (with glutamine and HEPES; Invitrogen) supplemented with: 75 ml heat-inactivated fetal bovine serum (FBS; Invitrogen) 5 ml penicillin (10,000 U/ml)/streptomycin (10,000 μg/ml); Invitrogen 5 ml MEM nonessential amino acids (100×; Invitrogen) 0.5 ml β-mercaptoethanol (0.7 mM; Sigma-Aldrich) Leukemia inhibitor factor (LIF): add 1000 U/μl LIF (Millipore) to a final concentration of 1 U/μl to mESC cultures just before use continued

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Store (without LIF) at 2◦ to 6◦ C for no longer than 4 weeks! Each batch of FBS is tested for optimal culture expansion and differentiation of mESCs. LIF is not stable at 37◦ C and therefore is stored separately at 4◦ C.

mESC differentiation medium (500 ml) 390 ml IMDM (with glutamine and HEPES; Invitrogen) supplemented with: 100 ml heat-inactivated fetal bovine serum (FBS; Invitrogen) 5 ml penicillin (10,000 U/ml)/streptomycin (10,000 μg/ml); Invitrogen 5 ml MEM nonessential amino acids (100×; Invitrogen) 0.5 ml β-mercaptoethanol (0.7 mM; Sigma-Aldrich) Store at 2◦ to 6◦ C for no longer than 4 weeks! COMMENTARY Background Information Isolation of cardiomyocytes from mouse embryos is limited due to relatively low cell numbers, particularly at early stages of development. In addition, culturing of adult cardiomyocytes is impossible, as the cells de-differentiate within days. At present, commercially available cardiac cell lines are rather specialized in cell type and developmental stage and do not recapitulate the physiological properties of cardiomyocytes. Pluripotent mESCs can serve as a valuable cell source of embryonic/perinatal cardiomyocytes. In vitro differentiation of mESCs closely resembles embryonic development, which allows for large-scale isolation of cells of all three germ layers, including specialized muscle cells of the heart at different stages of development (Doetschman et al., 1985; Robbins et al., 1990). For these reasons, in vitro differentiation of mESCs is a powerful tool to study cardiomyocyte commitment and differentiation under defined conditions. Moreover, the in vitro differentiation approach presented herein can be also used for human ESCs (UNIT 1D.2) or iPS cells (Kuzmenkin et al., 2009). In particular, the latter enables the generation of in vitro models for cardiovascular disease using patients as a source for iPS cells. Another advantage of mESCs is their straightforward genetic manipulation; singlecell clones can be selected and grown to obtain genetically homogeneous cell populations. With such an approach, it is, for example, possible to enrich a pure population of cardiomyocytes by using an antibiotic resistance gene under the control of a cardiacspecific promoter (Klug et al., 1996; Kolossov et al., 2006). Alternatively, cardiomyocytes can be strongly enriched (>95%) by FACS using live reporter gene expression under the control of cardiac promoters. Similarly, cardiovascular progenitor cells can be harvested

from EBs using early mesodermal markers such as Brachyury-T and Flk-1 (Kattman et al., 2006) in combination with live reporter genes such as GFP or RFP (green/red fluorescent protein). This allows for the isolation and expansion of proliferating cells to obtain higher yields of cardiomyocytes. Although the authors did not gain beneficial effects using the following additives, various medium supplements are reported to induce or enhance differentiation towards the mesodermal lineage and/or cardiomyocytes. The most promising substances reported to date are bone morphogenic proteins (BMP2, BMP4), transforming growth factor β(TGF-β), activin A, retinoic acid, platelet-derived growth factor BB (PDGF-BB), or sphingosine-1-phosphate (SPP) (Wobus et al., 1997; Sachinidis et al., 2003; Singla and Sobel, 2005; Laflamme et al., 2007; Puceat, 2008).

Critical Parameters and Troubleshooting Cell maintenance High percentage of dead cells after passaging: mESCs are quite sensitive to mechanical stress. Thus, slow and gentle pipetting is required. Colonies look dark/flat under phasecontrast microscope (Fig. 1F.11.3C): Make sure that the seeded cells are spread evenly over the tissue culture surface so that the colonies do not touch each other (see Fig. 1F.11.3B), because this may induce differentiation of the mESCs and loss of pluripotency and self-renewal capability. Cells should be seeded at a density that allows for passaging every second to third day. It is important to note that growth rate significantly varies between mESC lines/clones, which can largely be compensated by plating density. Therefore, new cultures need to be closely monitored and

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plating density for each cell line/clone has to be adjusted by trial and error. Yield of cardiomyocytes declines for a specific mESC line/clone over passages: mESCs can be maintained in culture indefinitely under appropriate conditions. However, the authors have found that prolonged culture of mESCs decreases their differentiation capacity toward the cardiac lineage after 25 to 30 passages, depending on the specific line/clone. Therefore, it is recommended to maximally expand early passages and freeze as many early-passage cells as possible. They should be stored as master stocks in liquid nitrogen and thawed only when cells of later passages show signs of senescence, e.g., when cardiomyocyte yield after differentiation decreases. Cell differentiation Condensed water on the lids of suspension culture dishes: This is due to the heat emanating from the horizontal shaker. It indicates that temperature in the culture dish rose higher than standard conditions, which may alter differentiation characteristics. In order to prevent this occurrence, place two to three empty dishes beneath the dishes containing EBs to buffer the temperature rise. EBs in suspension culture are not roundshaped: Incubate the EBs at lower density. Additionally, modify shaking velocity to lower the frequency of EBs contacting each other. Contact may lead to fusion of EBs resulting in larger aggregates with irregular shape and altered differentiation characteristics. Cardiogenic differentiation efficiency is variable: Keep constant culturing conditions especially on the first days of differentiation when EBs form. Fluctuations in CO2 concentration and temperature due to frequent opening of the incubator may hamper the cardiogenic differentiation of mESCs. Using the mass culture protocol, stoppage of the horizontal shaker must be prevented during the first 3 days of differentiation.

Anticipated Results

Differentiation of Mouse ESCs into Cardiomyocytes

These protocols describe the differentiation of mESCs in cell aggregates (EBs). The number of cells per EB at a given time point in differentiation can be roughly estimated from experience. On day 2 of differentiation, every EB will contain between 2000 and 3000 cells and will grow at a rate of about 1000 cells per day up to day 10. The yield of cardiomyocytes in the total cell population is rather low (less than 5%). From experience, the number of cardiomyocytes obtained using the hanging-drop

protocol is about 300 per EB at day 10 of differentiation; with the mass culture technique it will be roughly half that number. Nevertheless, due to the easier handling, the initial number of EBs obtained by the mass culture is much higher than with the hanging-drop method, leading to an overall increased cell number. Thus, although the fraction of cardiomyocytes per EB is lower, the overall yield is increased. Cardiomyocyte yield may be increased by various supplements as well as selection (see Background Information). With the increasing size of EBs, the diffusion of oxygen and nutrients into the center of the EB is reduced, resulting in apoptosis and also necrosis in the core areas of EBs. This, together with reduced proliferation rate, accounts for the fact that the cell number per EB does not further increase at later stages of differentiation. Plating of EBs can alleviate this problem, since they spread and flatten out, which in turn eases diffusion. Cardiomyocyte proliferation rapidly decreases after day 7 to 8 of differentiation, the time point when beating areas become visible. At day 9 of differentiation only ∼2% of ESC-derived cardiomyocytes are proliferating; at day 12, cardiomyocyte proliferation is almost completely abolished (Kolossov et al., 2006). The cardiomyocytes obtained with these in vitro differentiation methods resemble all stages of fetal development, as shown by genetic and electrophysiological studies (Hescheler et al., 1997). EBs are dissociated at day 9 to 11 of differentiation to harvest cardiomyocytes at an early developmental stage (roughly equivalent to mouse embryo day 9 to 12); later developmental stages are harvested at day 16 to 19 of differentiation (roughly equivalent to mouse embryo day 15 to 18). Notably, mESC-derived cardiomyocytes are not terminally differentiated, although plating of the cells induces further maturation shown by a pronounced cross-striation in immunocytochemistry (ICC) and at the molecular level (PCR, immunoblot; Hescheler et al., 1997; Boheler et al., 2002). Most of the data suggest that late stage mESC-derived cardiomyocytes resemble late fetal cardiomyocytes of mouse; similarly, human ESC-derived cardiomyocytes are rather immature cells (Snir et al., 2003; Sartiani et al., 2007). Examples of markers used to assess cardiogenic differentiation in more detail are the early mesodermal markers such as BrachyuryT, Mesp-1, Flk-1, early cardiac markers (transcription factors) like GATA-4, Nkx2.5, Isl-1, MEF2c, Tbx-5, and later cardiac markers

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(structural proteins) like α/β-MHC, α-actinin, c-troponin T/I, and MLC-2v.

Time Considerations Preparation of feeder flasks should be carried out at least 1 day before culturing mESCs. Frozen mESCs are cultured for 2 to 4 days and should be passaged at least one time (three passages are recommended) before starting differentiation (2 to 3 days per passage). Cardiac cells are first seen at day 7 to 8 of differentiation. Thus, the minimum time required to generate early-stage cardiomyocytes from frozen mESCs is about 2 weeks, or with the recommended passaging prior to differentiation, about 3 weeks. Generation of late stage cardiomyocytes accordingly prolongs the cultivation period by about 1 week.

Acknowledgement This work was supported by the European Commission FP7 Grant 223372 CardioCell Consortium.

Literature Cited Boheler, K.R., Czyz, J., Tweedie, D., Yang, H.T., Anisimov, S.V., and Wobus, A.M. 2002. Differentiation of pluripotent embryonic stem cells into cardiomyocytes. Circ. Res. 91:189-201. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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. Hescheler, J., Fleischmann, B.K., Lentini, S., Maltsev, V.A., Rohwedel, J., Wobus, A.M., and Addicks, K. 1997. Embryonic stem cells: A model to study structural and functional properties in cardiomyogenesis. Cardiovasc. Res. 36:149-162. Kattman, S.J., Huber, T.L., and Keller, G.M. 2006. Multipotent flk-1+ cardiovascular progenitor cells give rise to the cardiomyocyte, endothelial, and vascular smooth muscle lineages. Dev. Cell 11:723-732. Klug, M.G., Soonpaa, M.H., Koh, G.Y., and Field, L.J. 1996. Genetically selected cardiomyocytes from differentiating embryonic stem cells form stable intracardiac grafts. J. Clin. Invest. 98:216224. Kolossov, E., Bostani, T., Roell, W., Breitbach, M., Pillekamp, F., Nygren, J.M., Sasse, P., Rubenchik, O., Fries, J.W., Wenzel, D., Geisen, C., Xia, Y., Lu, Z., Duan, Y., Kettenhofen, R., Jovinge, S., Bloch, W., Bohlen, H., Welz, A., Hescheler, J., Jacobsen, S.E., and Fleischmann, B.K. 2006. Engraftment of engineered ES cellderived cardiomyocytes but not BM cells restores contractile function to the infarcted myocardium. J. Exp. Med. 203:2315-2327.

Kuzmenkin, A., Liang, H., Xu, G., Pfannkuche, K., Eichhorn, H., Fatima, A., Luo, H., Saric, T., Wernig, M., Jaenisch, R., and Hescheler, J. 2009. Functional characterization of cardiomyocytes derived from murine induced pluripotent stem cells in vitro. FASEB J. 23:4168-4180. Laflamme, M.A., Chen, K.Y., Naumova, A.V., Muskheli, V., Fugate, J.A., Dupras, S.K., Reinecke, H., Xu, C., Hassanipour, M., Police, S., O’Sullivan, C., Collins, L., Chen, Y., Minami, E., Gill, E.A., Ueno, S., Yuan, C., Gold, J., and Murry, C.E. 2007. Cardiomyocytes derived from human embryonic stem cells in pro-survival factors enhance function of infarcted rat hearts. Nat. Biotechnol. 25:10151024. Nagy, A., Rossant, J., Nagy, R., Bramow-Newerly, W., and Roder, J.C. 1993. Derivation of completely cell culture-derived mice from earlypassage embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 90:8424-8428. Puceat, M. 2008. Protocols for cardiac differentiation of embryonic stem cells. Methods 45:168171. Robbins, J., Gulick, J., Sanchez, A., Howles, P., and Doetschman, T. 1990. Mouse embryonic stem cells express the cardiac myosin heavy chain genes during development in vitro. J. Biol. Chem. 265:11905-11909. Sachinidis, A., Fleischmann, B.K., Kolossov, E., Wartenberg, M., Sauer, H., and Hescheler, J. 2003. Cardiac specific differentiation of mouse embryonic stem cells. Cardiovasc. Res. 58:278291. Sartiani, L., Bettiol, E., Stillitano, F., Mugelli, A., Cerbai, E., and Jaconi, M.E. 2007. Developmental changes in cardiomyocytes differentiated from human embryonic stem cells: A molecular and electrophysiological approach. Stem Cells 25:1136-1144. Singla, D.K. and Sobel, B.E. 2005. Enhancement by growth factors of cardiac myocyte differentiation from embryonic stem cells: A promising foundation for cardiac regeneration. Biochem. Biophys. Res. Commun. 335:637-642. Snir, M., Kehat. I., Gepstein. A., Coleman. R., Itskovitz-Eldor. J., Livne, E., and Gepstein, L. 2003. Assessment of the ultrastructural and proliferative properties of human embryonic stem cell-derived cardiomyocytes. Am J. Physiol. Heart Circ. Physiol. 285:H2355-H2363. Wobus, A.M., Wallukat, G., and Hescheler, J. 1991. Pluripotent mouse embryonic stem cells are able to differentiate into cardiomyocytes expressing chronotropic responses to adrenergic and cholinergic agents and Ca2+ channel blockers. Differentiation 48:173-182. Wobus, A.M., Kaomei, G., Shan, J., Wellner, M.C., Rohwedel, J., Ji, G., Fleischmann, B., Katus, H.A., Hescheler, J., and Franz, W.M. 1997. Retinoic acid accelerates embryonic stem cellderived cardiac differentiation and enhances development of ventricular cardiomyocytes. J. Mol. Cell Cardiol. 29:1525-1539.

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The Differentiation of Distal Lung Epithelium from Embryonic Stem Cells

UNIT 1G.1

Benjamin E. Van Vranken,1 Helen J. Rippon,2 Ali Samadikuchaksaraei,3 Alan O. Trounson,1 and Anne E. Bishop2 1

Monash University, Clayton, Australia Imperial College London, London, England 3 Iran University of Medical Sciences, Tehran, Iran 2

ABSTRACT The potential for embryonic stem (ES) cells to differentiate into cells with a distal lung epithelial phenotype has been demonstrated using different in vitro culture methods. Three separate protocols are described here that utilize both murine and human ES cells. The distal lung epithelial phenotype is induced through the use of embryonic distal lung mesenchyme in coculture systems with differentiating embryoid bodies or the use of soluble factors in defined media to maximize definitive endoderm formation and select and maintain the desired phenotype. Phenotypic analysis is demonstrated using immunocytochemistry and SP-C promoter-eGFP reporter gene expression in transgenic ES cells. These methods provide an increased efficiency of distal lung epithelial derivation from ES cells and, therefore, they provide the foundation for the development of a cell replacement product to treat chronic lung disease or a useful in vitro model for the study C of lung disease and development. Curr. Protoc. Stem Cell Biol. 2:1G.1.1-1G.1.22.
2007 by John Wiley & Sons, Inc. Keywords: embryonic stem cell r coculture r distal lung epithelium r differentiation

INTRODUCTION The protocols described in this unit are used for the differentiation of embryonic stem (ES) cells into epithelial progenitors of the distal lung. Embryonic stem cells have the potential to differentiate into derivates of the three embryonic germ layers as well as some extra-embryonic components. The authors’ research into the directed differentiation of ES cells into lung epithelial phenotypes has resulted in a number of different methods that can be used to accomplish this objective. The ultimate aim of the work is to derive these cells from ES cell cultures and use them to replace diseased or damaged distal lung epithelium, such as that which occurs in idiopathic pulmonary fibrosis or cystic fibrosis. Alternatively, the derived lung epithelium could be used to better understand lung development or the pathophysiology of lung disease—i.e., as a tool in drug discovery. Three protocols are described here; the two more complex methods have been validated on murine ES cells and the third has been shown to be effective for human and murine cells. Murine ES cells have been encouraged to differentiate into distal lung epithelial progenitors by coculturing them with murine distal embryonic lung mesenchyme (Basic Protocol 1) or by exposing them to different growth media formulations in a specific sequence (Basic Protocol 2). Human ES cells have been differentiated into the target phenotype using a defined growth medium (Basic Protocol 3). NOTE: For all procedures described in this unit, facilities for tissue culture, reagent preparation, wash-up, and sterilization are required. Experiments should be performed under sterile conditions in either Class II Biological Hazard Flow Hoods or laminar flow horizontal draft hoods. When working with human ES cells, Class II Biological Hazard Flow Hoods are recommended. Current Protocols in Stem Cell Biology 1G.1.1-1G.1.22 Published online July 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01g01s2 C 2007 John Wiley & Sons, Inc. Copyright


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NOTE: Ethical approval for the described protocols is required from the appropriate institutional research office. BASIC PROTOCOL 1

COCULTURE OF DIFFERENTIATING MURINE EMBRYOID BODIES WITH MURINE DISTAL EMBRYONIC LUNG MESENCHYME Embryoid body (EB) formation is a common technique used to initiate ES cell differentiation. These structures are thought to recapitulate the early developmental patterns of the embryo, as evidenced by their similarities in gene and protein expression patterns (Keller, 1995). However, EBs lack the spatial organization that occurs as a result of axis formation, such as that seen in the normal murine embryo (Doetschman et al., 1985). If allowed to differentiate spontaneously in suspension culture, EBs express mRNA and protein markers associated with definitive endoderm formation in the murine embryo (Van Vranken et al., 2005). By combining the differentiated EBs with murine embryonic distal lung mesenchyme, it is possible to direct the differentiation of the local ES cell population to a distal lung epithelial phenotype. This protocol is an adaptation of one developed by John Shannon (Shannon, 1994). In that article, the ability of embryonic distal lung mesenchyme to induce the distal lung epithelial phenotype in proximal lung epithelium was demonstrated. Here, the murine embryonic distal lung mesenchyme pieces, separated from the epithelium as described, are cultured in direct contact with EBs that are allowed to differentiate spontaneously for 8 to 10 days. This allows for the shortest possible distance, and therefore the highest concentrations, for soluble factors to be exchanged between the two tissues, and also allows for any signaling that is mediated by the extracellular matrix proteins or cell-cell signaling. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

Materials

Differentiation of Distal Lung Epithelium

Undifferentiated murine ES cell culture (feeder-free;) in gelatinized (see recipe) 25-cm2 tissue culture flask Phosphate-buffered saline (PBS; Invitrogen, cat. no. 20012-027) 0.025% (w/v) trypsin/EDTA (see recipe) 1× embryoid body differentiation medium (1× EBDM; see recipe) Pregnant BALB/c mice time-mated to embryonic day 12.5 70% (v/v) ethanol Hank’s Balanced Salt Solution (HBSS; Invitrogen) containing 1× antibiotic-antimycotic (AA; penicillin/streptomycin/amphotericin B; obtain 100× stock from Invitrogen) 50 mg/ml dispase (Sigma) in HBSS Coculture substratum (see recipe) in 3.5-cm diameter petri dishes PicoPure RNA Isolation Kit (Arcturus) Appropriate primers for phenotyping by RT-PCR (see recipe) 4% (w/v) paraformaldehyde in PBS Rat tail collagen gel, neutralized (Support Protocol) 50%, 70%, 90%, 95%, and 100% (v/v) ethanol Xylene Paraffin Appropriate probes for phenotyping by immunohistochemistry: e.g., anti-pro-SP-C antibody (Chemicon, cat. no. AB3786) or anti-TTF-1 antibody (NovoCastra, http://www.vision-bio.com, cat. no. NCL-TTF-1)

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15- and 50-ml conical centrifuge tubes (e.g., BD Falcon) Centrifuge 10-cm diameter bacteriological-grade petri dishes (BD Falcon) Dissecting instruments (soaked in 70% ethanol and rinsed in sterile PBS prior to use): Dissecting scissors, sharp Dissecting forceps Micro-dissecting scissors (Fine Science Tools) Fine-tipped dissecting forceps (Fine Science Tools) Dissecting microscope Glass embryo dish (Canemco & Marivac; http://www.canemco.com) 125-µm tungsten needles and needle holders (Fine Science Tools) Finely drawn glass Pasteur pipets and rubber bulb Embedding cassette Automated embedding machine Microtome Additional reagents and equipment for cell culture techniques including counting cells (Phelan, 2006), euthanasia of the mouse (Donovan and Brown, 2006), reverse transcription–polymerase chain reaction (RT-PCR; Beverley, 2001), preparation for immunohistochemistry including paraffin embedding (Zeller, 1989), and immunohistochemistry (Watkins, 1989) NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Culture cells for embryoid body formation 1. Aspirate medium from tissue culture flask containing feeder-free undifferentiated murine ES cells when subconfluent (i.e., 70% to 80% confluent). 2. Rinse growing surface with 4 ml PBS at room temperature. 3. Add 2 ml of 0.025% (v/v) trypsin/EDTA and incubate 2 to 3 min. at 37◦ C. 4. Inactivate trypsin by adding 4 ml of 1× EBDM down the growing surface to collect all cells. 5. Transfer cells and medium to a 15-ml centrifuge tube and centrifuge 5 min at ∼500 × g, room temperature. 6. Aspirate the supernatant and resuspend the cell pellet in 1 ml 1× EBDM. Count cells using a hemacytometer (Phelan, 2006). 7. Pipet 15 ml of 1× EBDM into each of the desired number of 10-cm bacteriologicalgrade petri dishes and transfer the appropriate volume of the cell suspension to seed 2.5 × 106 cells in each dish. Return dishes to incubator. Bacteriological-grade plastic should prevent the derived embryoid bodies (EBs) from adhering to the surface. If the EBs do adhere, it is likely that the ES cell cultures from which they were derived were too differentiated. It is a good idea to test these cultures regularly for the undifferentiated phenotype by immunostaining for markers such as Oct-4, Nanog, and Sox2 (Sun et al., 2006). EBs allowed to differentiate spontaneously in serum-containing medium using this method were found to express GATA-4 and Foxa2 mRNA and proteins, both necessary for definitive endoderm formation, after 8 to 10 days in culture (Leahy et al., 1999; Van Vranken et al., 2005). Current Protocols in Stem Cell Biology

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8. Change the medium in these cultures every 2 to 3 days. This can be done by allowing the EBs to settle by gravity at room temperature, but it is quicker to transfer them to a 15-ml centrifuge tube and gently centrifuge them for 2 min at 10 to 15 × g, room temperature. The supernatant can then be carefully removed and fresh 1× EBDM used to resuspend the EBs. Care should be taken not to pipet the EBs up and down too much, as this will disaggregate them.

Isolate murine embryo 9. Sacrifice the desired number of pregnant BALB/c female mice, time-mated to embryonic day 12.5 (E12.5), by cervical dislocation (Donovan and Brown, 2006). Euthanasia should be performed according to ethical approval obtained from the Institutional Animal Care and Use Committee. Alternatives to BALB/c mice should be considered if this strain does not yield reliable numbers of embryos. This mouse strain can yield anywhere from two to ten embryos per pregnant female.

10. Turn the mouse onto its back and spray the abdomen with 70% ethanol. Using sharp dissecting scissors and forceps, lift up the abdominal wall and make a mid-sagittal incision from pubis to xiphisternum. Transverse cuts across the lower abdomen will allow for better visualization of the uterus.

11. Locate the two horns of the murine uterus, lateral to the intestines. Grasp the upper pole with a forceps and lift each horn containing the embryos. Cut the upper and lower poles and transfer the uterus to a 50-ml tube containing ice-cold HBSS supplemented with 1× AA. The mouse carcass should be disposed of according to institutional guidelines.

12. Transfer the uterus to a 10-cm bacteriological-grade petri dish containing 10 ml ice-cold HBSS supplemented with 1× AA, placed on the stage of a dissecting microscope. The dissecting microscope should be placed inside a laminar flow cabinet to reduce the likelihood of infections occurring in the cultures.

Dissect lung tissue 13. Using micro-dissecting scissors and fine forceps, grasp each embryo and cut a hole, large enough for the embryo to exit from, in the wall of the uterus and through the amniotic sac. The embryo should be freed from the amniotic sac, leaving the placenta behind within the uterus.

14. Cut the umbilical cord of the embryo with the micro-dissecting scissors. 15. Grasp the abdomen of the embryo with the forceps and remove its head with the scissors. 16. Make a mid-sagittal incision through the anterior chest wall, followed by two transverse cuts below the ribs. 17. Grasp the heart, lungs, and esophagus with the forceps and lift them as a unit out of the chest cavity. The lower end of the esophagus just near the stomach should be visible, as it is tethered to the contents of the chest cavity. Differentiation of Distal Lung Epithelium

18. Cut the lower end of the esophagus, freeing it from the rest of the embryo.

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Figure 1G.1.1 A pair of embryonic lungs after removal from a gestational day 12.5 (E12.5) embryo. Note the distal epithelial rudiments and associated mesenchyme. Original magnification, 100×.

19. Transfer the heart, lungs, and esophagus to a clean part of the dish. Carefully separate the embryonic lung (Fig. 1G.1.1) from the other tissues. Transfer the lung to a sterile 1.5-ml microcentrifuge tube using a micropipet tip and place on ice. When transferring tissues, be sure first to coat the inside of the micropipet tip or glass Pasteur pipet with 1× EBDM. The tissues tend to stick to the inside of the pipet if this is not done. The embryonic lung is ∼1 mm long and an indefinite number can be placed in the same microcentrifuge tube.

20. Repeat steps 9 to 19 for all mice and embryos.

Separate the distal embryonic lung mesenchyme 21. Transfer the embryonic lungs to a glass embryo dish containing 2 ml of ice-cold HBSS supplemented with 1× AA, placed on the stage of the dissecting microscope. 22. Using two tungsten needles, in a scissors-like motion, cut the distal lung buds from the rest of the embryonic lung, leaving the trachea and most of the proximal airways (Fig. 1G.1.1). It may be easier to use one tungsten needle and a 22-G syringe needle to remove the lung buds. The goal is to remove the distal lung tips that are actively budding from the bronchial tree. A single embryonic lung may yield up to ten distal tips, depending on the age. This will be enough to perform two to three direct-contact cocultures.

23. Discard the trachea and proximal lung components (unless using this material as a negative control). 24. Carefully aspirate the HBSS/AA without removing the distal lung tips. 25. Add 2 to 3 ml of 50 mg/ml dispase in HBSS to the embryo dish and incubate at 37◦ C for 15 to 20 min. The dispase treatment separates the two layers (epithelial rudiments and mesenchyme) without disaggregating the cells within the layers.

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26. Aspirate dispase and replace with HBSS supplemented with 1× AA. Aspirate the wash solution and add fresh HBSS supplemented with 1× AA. Repeat this washing two to three times. Dispase cannot be inactivated with serum, so it is necessary to wash residual dispase away from the distal tips.

27. Using the tungsten needles, carefully tease the mesenchyme away from the epithelial rudiment. Transfer the mesenchyme pieces to a sterile 1.5-ml microcentrifuge tube and place on ice. This process may be aided by the addition of lyophilized DNase. Dip the tungsten needle into the powder, in order to pick up a few particles. Transfer these back to the embryo dish and mix. DNA released from cell debris makes the tissue clump together and stick to the needles. Take care to remove as much epithelium as possible in order to prevent the contamination of the cocultures.

Establish direct contact coculture of EBs with murine embryonic distal lung mesenchyme A positive control consisting of distal lung mesenchyme (prepared as described above) plus microdissected distal lung endoderm (Shannon et al., 1994; Van Vranken et al., 2005) is recommended, as well as negative controls consisting of EBs alone, EBs plus microdissected tracheal lung mesenchyme, and EBs plus microdissected gut mesenchyme (Van Vranken et al., 2005). The preparation procedures for these tissues are as described in the preceding steps of this protocol. 28. For each coculture, randomly choose an EB that has been allowed to differentiate spontaneously (see steps 7 to 8) for 8 to 10 days and transfer it to a coculture substratum in a 3.5-cm petri dish using a glass Pasteur pipet and rubber bulb. Three to five cocultures may be done on the same substratum in the same dish under the same culture conditions and in the same medium.

29. Transfer three to six pieces of distal lung mesenchyme (from step 27) to the substratum in the vicinity of each EB. Place the dish on the stage of the dissecting microscope. While watching carefully through the microscope, slowly remove excess medium. 30. Using a tungsten needle, gently position the EB with the mesenchyme pieces nearly surrounding it. 31. With the tip of the finely drawn glass Pasteur pipet, gently make indentations in the substratum around the EB and mesenchyme pieces, to form a sort of “moat” around the tissue pieces. 32. Use the tungsten needle to position the mesenchyme pieces closer to the EB, so that they are in direct contact. 33. Add a few small drops of 1× EBDM to the “moat” and near the coculture using the finely drawn glass Pasteur pipet, being careful not to disrupt the tissues. Too much medium will cause the pieces of tissue to float away. Too little will allow the tissue to dry out.

34. Incubate the cocultures overnight at 37◦ C in 5% CO2 . Differentiation of Distal Lung Epithelium

After the initial overnight incubation, the mesenchyme-EB interface should still be observed around the outside of the EB (Fig. 1G.1.2). After 48 hr, the two tissues will fuse together into one tissue construct.

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Figure 1G.1.2 200×.

A direct-contact coculture after initial 24 hr of incubation. Original magnification,

35. On the following day, add 200 µl of 1× EBDM near the edge of the dish, taking care not to disrupt the tissues. 36. Culture the tissue constructs for 5 or 12 days, adding fresh medium or changing the medium every 2 to 3 days. The tissue constructs will float near the surface of the medium, providing an air-liquid interface, which is thought to maintain the type II phenotype (Alcorn et al., 1997; Xu et al., 1998).

Phenotype the cocultured tissue constructs To phenotype by RT-PCR 37a. At the end of the coculture period, isolate total RNA using the PicoPure RNA Isolation Kit. 38a. Perform reverse transcription–polymerase chain reaction (RT-PCR; Beverley, 2001) using the appropriate primers for phenotyping: e.g., surfactant protein C (SP-C) forward and reverse primers (see Reagents and Solutions).

To phenotype by immunohistochemistry 37b. Fix tissues in 4% paraformaldehyde in PBS for 20 min at room temperature. 38b. Embed tissue in neutralized rat tail collagen gel. Before paraffin embedding, the tissue construct is much easier to handle if it is first embedded in a neutralized collagen gel. The neutralized collagen solution (prepared as described in the Support Protocol ) is liquid at 4o C, but solidifies at room temperature. Thus, the tissue construct can be enveloped (“enrobed”) in cold neutralized collagen gel solution (in other words, the sample is mixed with the gel solution, but not so much that the tissue disaggregates), which is then allowed to solidify at room temperature.

39b. Place the collagen gel, with tissue construct contained within it, into an embedding cassette and use an abbreviated embedding technique in an automated tissue embedding machine (30 min each in ascending concentrations of ethanol: 50%, 70%, Embryonic and Extraembryonic Stem Cells

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90%, 95%, twice in 100%), then twice for 1 hr in xylene and twice for 1 hr in molten paraffin). Paraffin embedding of tissue is also described in Zeller (1989).

40b. Section the paraffin block using a microtome (also see Zeller, 1989). Perform tissue staining (Watkins, 1989) using the appropriate antibody probes for phenotyping: e.g., anti–prosurfactant protein C antibody (anti-pro-SP-C) or anti–thyroid transcription factor 1 antibody (anti-TTF-1). SUPPORT PROTOCOL

PREPARATION OF NEUTRALIZED RAT-TAIL COLLAGEN GEL Neutralized rat tail collagen gel is used to “enrobe” tissues before paraffin embedding (see Basic Protocol 1). This allows for easier handling of samples during the paraffinembedding process.

Materials Rat tail collagen (Sigma, cat. no. C-8897) 0.1% (v/v) acetic acid Medium 199 (Sigma) 0.35 M NaOH (Sigma) Centrifuge with Beckman JA-14 rotor (or equivalent) 1. Place 100 mg rat tail collagen in 15 ml of 0.1% (v/v) acetic acid. Extract soluble collagen over a 48-hr period in a continuously agitated container at 4◦ C. 2. Centrifuge the collagen solution 20 min at 3800 × g, 4◦ C, to remove debris. Remove supernatant and store up to 1 year at 4◦ C. 3. When ready to use, mix 1.8 ml of the ice-cold collagen solution with 0.2 ml medium 199. Neutralize by adding ∼98 µl of 0.35 M NaOH. Keep the mixture on ice throughout the procedure. The color of the resulting solution must be red-orange in order for the gel to set properly. The neutralized gel can be used to embed a tissue sample (see Basic Protocol 1, step 38b), which will solidify after 15 to 20 min at room temperature. ALTERNATE PROTOCOL

INDIRECT COCULTURE OF MURINE EBs AND MURINE EMBRYONIC DISTAL LUNG MESENCHYME This protocol can be used to examine the effects of soluble factors secreted by the mesenchyme without the influence of cell-cell signaling or some extracellular matrix proteins, which takes effect when the two tissues are in direct contact. The EBs are separated from the mesenchyme by a 0.02-µm-pore Anapore membrane, which does not permit cell contact or migration. Soluble factors secreted by the mesenchyme, however, can diffuse across the membrane and influence the differentiation of the EBs.

Additional Materials (also see Basic Protocol 1) 24-well tissue culture plates 0.02-µm Anapore membrane tissue culture inserts for 24-well plates (Whatman) NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. Differentiation of Distal Lung Epithelium

NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

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1. Perform steps 1 to 27 of Basic Protocol 1. After separation from the epithelium (Basic Protocol 1, step 27), transfer the murine embryonic distal lung mesenchyme pieces from two lungs to a well of a 24-well plate. The exact amount of mesenchyme needed is unknown. In combination with the EBs, the mesenchyme from two lungs was determined to be the maximum amount of tissue that could be used without the medium getting too acidic between daily medium changes. This should theoretically result in the highest concentration of soluble factors in the medium. The inclusion of lung epithelium in these cultures may improve the expression of SP-C mRNA in the EBs after coculture.

2. Cover the mesenchyme with 1 ml 1× EBDM and place in the incubator at 37◦ C for 30 min to allow for attachment. 3. Using a sterile forceps, place a 0.02-µm Anapore membrane tissue culture insert into each of the appropriate wells. 4. Choose three EBs at random from an 8- to 10-day suspension culture. Transfer the EBs to each culture insert along with 200 µl of 1× EBDM. 5. Incubate the cocultures for 5 or 12 days at 37◦ C in 5% CO2 . Change the medium in both chambers (above and below the filter) every 1 to 2 days. The EBs should be maintained just near the air-liquid interface throughout the remaining incubation period, in an effort to maintain the type II pneumocyte phenotype (Alcorn et al., 1997; Xu et al., 1998) and allow for adequate gas exchange for both the EBs and mesenchyme. A positive control consisting of distal lung mesenchyme (prepared as described in Basic Protocol 1) plus microdissected distal lung epithelium is recommended, as well as negative controls consisting of EBs alone, EBs plus microdissected tracheal lung mesenchyme, and EBs plus microdissected gut mesenchyme. Preparation of these tissues is performed as described in Basic Protocol 1.

6. At the end of the coculture period, phenotype the tissue using RT-PCR as described in Basic Protocol 1, steps 37a to 38a.

DERIVATION OF EARLY DISTAL LUNG EPITHELIAL PROGENITORS FROM MURINE ES CELLS

BASIC PROTOCOL 2

Targeting ES cell differentiation to specific target cell phenotypes is a process of recapitulating natural embryonic development. As well as by coculture, this can be achieved through sequential manipulation of the cell culture medium in a manner designed to target and enhance the development of each cell type in the differentiation pathway in turn. The differentiation pathway for distal lung epithelial cells can be summarized as: ES cell/epiblast → mesendoderm → definitive endoderm → foregut endoderm → early distal lung progenitors → mature lung epithelium. This protocol employs a three-stage strategy to derive early distal lung epithelial progenitor cells thought to be representative of those present in the early branching lung at approximately E10 to E11 of murine development. This is achieved by the early application of high levels of activin A to improve the specification of mesendoderm and definitive endoderm, followed by two defined, serum-free media that allow the maturation of endoderm into lung progenitors.

Materials Undifferentiated murine ES cell culture (feeder-free) in gelatinized (see recipe) 25-cm2 tissue culture flask Definitive Endoderm-Inducing Medium (DEIM; see recipe) with and without activin A Phosphate-buffered saline (PBS; Invitrogen, cat. no. 20012-027) Small Airway Basal Medium (SABM; Cambrex)

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10-cm-diameter bacteriological petri dish 6-well tissue culture plate or 75-cm2 tissue culture flask, gelatinized (see recipe) Additional reagents and equipment for forming EBs (Basic Protocol 1, steps 1 to 7) NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Stage 1: 10-day suspension culture of embryoid bodies 1. Day 0: Form EBs as described in steps 1 to 7 of Basic Protocol 1. Culture EBs in suspension in 1× EBDM for 2.5 days. The authors have not observed significant differences in differentiation efficiency when using different cell seeding densities to initiate EB formation. The quality of the culture, however, is critical (see Critical Parameters and Troubleshooting).

2. Day 2.5: Transfer EBs into 10 ml of DEIM in a 10-cm-diameter bacteriological petri dish. Culture for 4.5 days at 37◦ C in 5% CO2 . Replace the medium on day 5. Definitive Endoderm-Inducing Medium (DEIM) is a serum-free medium containing saturating levels of recombinant activin A (100 ng/ml) Embryoid bodies are treated with high levels of activin A early in differentiation to enhance the formation of definitive endoderm, the germ layer from which lung epithelium is derived. The authors use only 10 ml of medium per dish during this stage, in order to economize on activin A.

3. Day 7: Transfer EBs into 15 ml DEIM without activin A. Continue to culture EBs in suspension for an additional 3 days.

Stage 2: 11-day adherent culture of embryoid bodies 4. Day 10: Distribute each petri dish of EBs between two gelatinized 6-well plates in fresh DEIM without activin A at 2 to 3 ml/well and incubate to allow outgrowth and maturation of the early differentiated murine ES cells. Alternatively, the EBs can be plated in gelatinized 75-cm2 tissue culture flasks in 10 ml DEIM without activin A. It is very important that EBs not be dissociated at this stage. Dissociation of EBs abolishes all lung epithelial differentiation, suggesting that the presence of three-dimensional structure is critical to the specification of lung epithelial cell types. The majority of EBs should adhere in the next 3 to 4 days; to encourage this process, do not disturb the plates more often than is absolutely necessary. EBs can be seeded at higher densities in fewer culture wells or flasks, but the authors’ experience is that this can negatively affect lung differentiation.

5. After most EBs have adhered, replace the medium twice weekly, removing any EBs that are still floating.

Stage 3: Selection of lung epithelial progenitors with lung-specific medium 6. Day 21: Aspirate medium and wash the cells with 2 ml PBS per well. Add 2 to 3 ml SABM to each well and culture for 4 to 10 days, replacing the medium at least twice weekly to remove dead cells. Differentiation of Distal Lung Epithelium

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7. At the end of the culture time, harvest the remaining live cells for analysis. The final stage selects for lung epithelial progenitors via the application of a serum-free commercial medium, SABM, which is optimized for the growth and survival of pulmonary epithelial phenotypes. This causes widespread cell death of non-lung epithelial cells, thus enriching the population for the desired cell type. Where cultures are intended for analysis by fluorescence methods, e.g., by flow cytometry, the extensive autofluorescence caused by this final step can entirely confound analysis. In the authors’ laboratory, this step is omitted if cells are destined for flow cytometry, fluorescence-activated cell sorting (FACS), or fluorescent immunocytochemistry. SABM has a short shelf life, and it is important that it be stored according to the manufacturer’s instructions. The authors recommend that SABM be used within 2 to 3 weeks of purchase.

DERIVATION OF DISTAL RESPIRATORY EPITHELIAL CELLS FROM HUMAN ES CELLS

BASIC PROTOCOL 3

Because the ultimate goal of the protocols described in this unit is to produce clinically applicable results, the authors have investigated the potential for the differentiation of human ES cells to be directed towards the distal airway epithelial phenotype. This experiment was modeled after an early protocol developed by the authors’ group for the differentiation of murine ES cells into lung epithelial cells (Ali et al., 2002). It was hypothesized that, if stem cells were cultured in a condition suitable for the maintenance of the primary distal lung epithelial cell phenotype, their differentiation into that phenotype would be enhanced. This culture condition can be provided by a defined medium called Small Airway Growth Medium (SAGM). The authors have shown that this medium can significantly enhance the differentiation of human ES cells into respiratory epithelial cells (Samadikuchaksaraei et al., 2006).

Materials Undifferentiated human ES cell (hESC) colonies on mitotically inactivated murine embryonic fibroblast feeder cells in 6-well plates Medium for maintaining undifferentiated hESC 1 mg/ml collagenase IV (see recipe) Phosphate-buffered saline (PBS; Invitrogen, cat. no. 20012-027) hEB medium (see recipe) Dif◦ hES medium (see recipe) Small Airway Growth Medium (SAGM; Cambrex) Felt-tip pen Inverted microscope Glass Pasteur pipet with tip broken, autoclaved 15-ml centrifuge tubes Centrifuge 90-mm cell adherence–resistant Nalgene polymethylpentene petri dishes (Nalge Nunc) 6-well tissue culture–treated plates with high-grade polystyrene culture surfaces (Orange Scientific; http:// www.orangesci.com) NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

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Stage 1: 7-day embryoid body culture 1. Day 0: If there are numerous differentiated colonies in addition to undifferentiated colonies in the hESC cultures, mark undifferentiated colonies with a felt-tip pen under the inverted microscope after identification. 2. About 1 hr before splitting, apply fresh medium to the cultures (same medium as is used to maintain the undifferentiated colonies). 3. Aspirate the medium from the culture plates and add sufficient 1 mg/ml collagenase IV to cover the culture surface. 4. Incubate the cells in the 37◦ C incubator for 3 to 5 min. 5. Add 3 ml of PBS to each well. Use the broken glass Pasteur pipet tip to scrape off and aspirate the undifferentiated hES cell colonies. 6. Transfer the undifferentiated cells to a 15-ml centrifuge tube and gently pipet them up and down to partially break up the large colonies. The cells must remain in clumps. If the colonies are broken into single cells, they will not proliferate.

7. Centrifuge the cells 5 min at 200 × g, room temperature. 8. Aspirate the supernatant and resuspend the cells with hEB medium. Transfer to 90-mm cell adherence–resistant Nalgene polymethylpentene petri dishes with a total of 15 ml hEB medium per dish. 9. Culture cell clumps in suspension in hEB medium in the cell adherence–resistant petri dishes for 7 days. Replace the medium every day.

Stage 2: Post-embryoid body culture 10. Day 7: Transfer EBs from the 90-mm petri dish into the wells of a 6-well tissue culture–treated plate. Incubate cells with 2.5 ml hEB medium per well. 11. Continue incubating for 2 days, replacing the medium every day. The EBs will adhere to the culture surfaces.

12. Day 9: Change to Dif◦ hES medium and continue incubating for 10 days, replacing the medium every day. 13. Day 19: Change the medium to SAGM and continue incubating for 5 days, replacing the medium every day. 14. Day 24: Harvest the cells for analysis by RT-PCR and immunophenotyping (see Basic Protocol 1).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized distilled water or equivalent for recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Coculture substratum Melt 1% (w/v) tissue culture–tested agarose (Sigma) prepared in tissue culture– tested distilled water in a microwave oven at high power for 1 to 2 min. Mix 1:1 with 2× EBDM (see recipe). Place medium in 3.5-cm-diameter petri dishes at 2 ml per dish and allow to cool at room temperature. Store up to 2 weeks at 4◦ C. Differentiation of Distal Lung Epithelium

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Collagenase IV solution, 1 mg/ml Dissolve collagenase IV powder in DMEM/F12 medium (Invitrogen) for a final concentration of 1 mg/ml. Filter sterilize using a 0.2-µm filter. Store up to 1 month at 4◦ C.

Definitive Endoderm-Inducing Medium (DEIM) High-glucose DMEM, without L-glutamine or sodium pyruvate, supplemented with: 10% (v/v) KnockOut Serum Replacement (KOSR; Invitrogen) 2 mM L-glutamine 100 ng/ml recombinant activin A (R&D Systems) Store up to 2 weeks at 4◦ C Differentiating hES cells (Dif ◦ hES) medium DMEM/F12 medium (Invitrogen) supplemented with: 20% (v/v) KnockOut Serum Replacement (Invitrogen) 2 mM L-glutamine 0.1 mM MEM nonessential amino acids (Invitrogen) 0.1 mM 2-mercaptoethanol (2-ME; Sigma) Store up to 2 weeks at 4◦ C Embryoid body differentiation medium (EBDM), 1× High glucose DMEM, without L-glutamine, or sodium pyruvate supplemented with: 10% (v/v) FBS 2 mM L-glutamine 1× penicillin/streptomycin solution (from 100× stock; Invitrogen) 0.1 mM 2-mercaptoethanol (2-ME; Sigma) Store up to 2 weeks at 4◦ C The FBS used for the propagation of undifferentiated ES cells will not necessarily give the best results in differentiation. A batch test should be performed on embryonic distal lung endoderm and mesenchyme cocultures to see which FBS batch best maintains the type II pneumocyte phenotype.

Embryoid body differentiation medium (EBDM), 2× (for preparing coculture substratum only) Empty the contents of one packet (makes 1 liter of 1 × DMEM) of powdered DMEM, high-glucose, without L-glutamine, sodium pyruvate, or sodium bicarbonate (Invitrogen) into a 500-ml clean, autoclaved glass bottle. Add 450 ml of tissue culture-tested distilled water and mix. Add the required amount (specified in package insert from Invitrogen) of sodium bicarbonate powder (Sigma) to medium and check pH. Adjust pH to ∼7.1 using sodium bicarbonate. Add tissue culture–tested distilled water up to 500 ml. Filter sterilize the medium using a vacuum-assisted 0.22-µm filter-sterilization unit (Nalgene). Add 7.6 ml of the medium to a 15-ml centrifuge tube. Add 2 ml of FBS, 0.2 ml of 100× L-glutamine (Invitrogen), and 0.2 ml of 100× penicillin/streptomycin (Invitrogen). Store up to 2 weeks at 4◦ C. The pH should increase by about 0.1 after filter sterilization.

Gelatinized tissue culture vessels Stock solution (1%): Prepare a 1% (w/v) gelatin stock by dissolving 0.4 g of gelatin powder (Sigma) in 40 ml tissue culture-tested distilled water. Autoclave, and store up to 2 months at −20◦ C. continued

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Working solution (0.1%): Dilute 5 ml of 1% (w/v) gelatin stock in 50 ml tissue culture–tested distilled water. Store up to 4 weeks at 4◦ C. Coating of vessels: Add the appropriate amount of 0.1% gelatin—1 ml (for wells of 6-well plate), 4 ml (for 25-cm2 tissue culture flask), or 12 ml (for 75-cm2 tissue culture flask). Incubate at room temperature for at least 15 min, then aspirate liquid.

hEB medium KnockOut D-MEM (Invitrogen) supplemented with: 20% (v/v) heat-inactivated fetal bovine serum (FBS, batch-tested for the cells) 2 mM L-glutamine 0.1 mM MEM nonessential amino acids (Invitrogen) Store up to 2 weeks at 4◦ C Primers for phenotyping by RT-PCR SP-C: SP-C1 (forward, exon 3-4 boundary): 5 -tatgactaccagcggctcct-3 SP-C3 (reverse, exon 5-6 boundary): 5 -gtttctaccgaccctgtgga-3 Primers SP-C1 and SP-C3 generate a 295-bp amplicon. β-actin: mβ-a1S (forward): 5 -gtcgtaccacaggcattgtgatgg-3 mβ-a1A (reverse): 5 -gcaatgcctgggtacatggtgg-3 The β-actin primers generate a 500-bp amplicon. HPRT (5 end): HPRT5 F (forward): 5 -actgctttccggagcggtag-3 HPRT5 R (reverse): 5 -gaacttatagcccccccttga-3 The HPRT (5 end) primers generate a 302-bp amplicon. HPRT (3 end): HPRT3 F (forward): 5 -tttgatttgcactatgagcctata-3 HPRT3 R (reverse): 5 -agcaaaacctcttagatgctgtta-3 The HPRT (3 end) primers generate a 272-bp amplicon. Trypsin/EDTA, 0.025% Dilute 20 ml of 0.05% (w/v) trypsin/50 mM EDTA (Invitrogen) in 20 ml PBS (PBS; Invitrogen, cat. no. 20012-027) for a final concentration of 0.025% (w/v). Store in aliquots up to 2 weeks at 4◦ C or up to 6 months at −20◦ C.

COMMENTARY Background Information

Differentiation of Distal Lung Epithelium

The derivation of specific differentiated cell types from embryonic stem (ES) cells may allow for breakthroughs in tissue engineering, drug discovery, and regenerative medicine. The use of these cells to study disease or in cell-replacement therapy could revolutionize science and medicine as we know it. Many chronic lung diseases, such as chronic obstructive pulmonary disease, or smoker’s lung, continue to result in the premature deaths of thousands of individuals around the world because they can only be cured by lung transplantation. Unfortunately, the supply of trans-

plantable lungs does not meet demand. Distal lung epithelium derived from ES cells may provide a ready supply of healthy cells, available to the clinician on demand. These cells, infused into damaged lung tissue, may also provide the health benefits without the morbidity associated with transplants of entire organs. Alternatively, the cells could be used to create in vitro models of disease. For example, lung epithelial progenitors derived from human ES cell lines that contain a cystic fibrosis transmembrane regulator mutation (Mateizel et al., 2006) could open up possibilities for basic science research and drug discovery. In

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vitro models of the distal lung are also in great demand as an animal-free approach to testing the respiratory toxicity of household and environmental chemicals. The authors of this unit have derived distal lung epithelium from ES cells using several different approaches. Originally, murine embryoid bodies (EBs) were cultured in serum-containing medium, and the small but unknown proportion of spontaneously differentiating cells were then selected in Small Airway Growth Medium (SAGM), a serum-free growth medium designed to maintain primary lung epithelial cells (Ali et al., 2002). This was followed by work which demonstrated that Small Airway Basal Medium (SABM), the basal medium on which SAGM is based (i.e., SAGM without the panel of added growth factors), actually resulted in a higher level of surfactant protein-C (SP-C) mRNA expression than SAGM itself (Rippon et al., 2004). During early attempts to direct lung epithelial differentiation, the authors of this unit recognized that the derivation of definitive endoderm is fairly inefficient in spontaneously differentiating cultures. The process by which definitive endoderm is specified in the gastrulating embryo is poorly understood. However, evidence from the field of developmental biology suggested that exposure to high levels of nodal signaling may be an important inductive cue (Lowe et al., 2001). Subsequently, activin A, a ligand for the nodal signaling pathway, was shown to induce the differentiation of a definitive endodermal-like population from undifferentiated murine ES cells (Kubo et al., 2004; Tada et al., 2005; Yasunaga et al., 2005). More recently, this has also been demonstrated using human ES cells, although it has been reported that concomitant suppression of phosphatidylinositol 3-kinase signaling is essential during activin A treatment to prevent the activation of self-renewal pathways instead of endoderm differentiation (D’Amour et al., 2005, 2006). The protocol for derivation of distal lung progenitors was further improved when activin A was utilized to induce the formation of definitive endoderm within EBs before treatment with SABM (Basic Protocol 2; Rippon et al., 2006). The use of serum-free media in most steps of Basic Protocol 2 allows for a more defined set of culture conditions, a critical factor in reproducibility. A significant proportion of cells (2% to 4%) in the resulting cultures were found to express SP-C mRNA. When considering that SP-C expression virtually never occurs within spon-

taneously differentiating cultures, this is a vast improvement. Human ES cells have also been induced to differentiate into distal epithelial cells by directing the differentiation of EBs using SAGM (Samadikuchaksaraei et al., 2006). The presence of the respiratory epithelial phenotype was confirmed by the detection of markers that are exclusively expressed in human lung epithelial cells, including surfactant proteins C (Phelps and Floros, 1991), A (Madsen et al., 2003), and B (Pryhuber, 1998; Strayer et al., 1998), and lamellar bodies (Voorhout et al., 1993). SAGM contains factors known to promote pneumocyte differentiation and/or function, such as epidermal growth factor, which promotes type II alveolar cell maturation (Plopper et al., 1992; Edwards et al., 1995), and retinoic acid, which has been found to significantly up-regulate the expression of surfactant protein genes by lung explants in vitro (Bogue et al., 1996). However, as with the protocols designed for differentiation of murine ES cells, the yield of the target cells with this protocol is low, and it needs further optimization. The coculture method (Basic Protocol 1 and Alternate Protocol) has been widely used. Here, a multitude of complicated cues required for the proper growth and phenotypic induction of another cell type are provided through coculture with a tissue that is known to provide those cues. This technique has been used to promote hematopoietic commitment (Nakano et al., 1996) and to derive dopaminergic neurons (Kawasaki et al., 2000), hepatocytes (Fair et al., 2003), and chondrocytes (Sui et al., 2003) from ES cells. In normal lung development, embryonic distal lung mesenchyme is thought to induce the formation of the type II pneumocyte phenotype through molecular cues in the form of signaling molecules, growth factors, cellcell interactions, and extracellular matrix proteins. Shannon (1994) demonstrated that, after 5 days in culture, it was possible for distal lung mesenchyme to induce the distal lung epithelial phenotype when grafted to proximal lung epithelium. This technique was performed by replacing lung epithelium with EBs yielding similar results. Epithelial tubules that were immunoreactive for TTF-1 and pro-SP-C proteins, the coexpression of which indicate the type II pneumocyte phenotype, were demonstrated in cocultures produced from EBs and murine embryonic distal lung mesenchyme (Van Vranken et al., 2005).

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Ultimately, the goal of any coculture experiment of this type is to determine which factors are required to produce the desired phenotype in the ES cells and replace the cocultured tissue with a combination of manufactured products, such as recombinant soluble factors and extracellular matrix proteins, which would recapitulate the effects of the tissue with the inductive capability. This reduces the number of unknown variables in the experiment introduced by the inducing tissue in the coculture model and would make the derived cell type eligible for transplantation into human beings. Cells that have been cultured with another cell population are not likely to be used to treat patients because of the possibility of undesirable elements in the finished product, e.g., viral particles or contaminating cells. However, until the exact culture conditions required to derive and maintain respiratory epithelial cells from ES cells have been elucidated, coculture remains a useful model for the derivation of these cells, not the least because this method results in branching of the derived epithelial structures within a three-dimensional tissue construct. Coculture is also the most efficient method that the authors have utilized to derive lung epithelium in terms of percentage yield, although it is also the most labor-intensive. Both direct-contact cocultures (Basic Protocol 1) and indirect cocultures (Alternate Protocol) may be used to direct the phenotype of differentiating ES cells. The indirect coculture method described in the Alternate Protocol can be used to direct the differentiation of the ES cells without the need for direct contact with the mesenchyme. The indirect coculture method may help to establish which soluble factors from the mesenchyme are sufficient to induce the lung epithelial phenotype in the ES cells. Antibodies raised against candidate soluble factors could abrogate induction in vitro and help to characterize the mechanism of induction. However, it is not known whether this method results in the production of TTF-1/proSP-C protein, which would verify the target cell phenotype.

Critical Parameters and Troubleshooting

Differentiation of Distal Lung Epithelium

General considerations As with all ES cell differentiation protocols, the quality of the original, undifferentiated population is critical. ES cells should exhibit good undifferentiated morphology prior to EB formation. In the case of murine ES cells, this is typically characterized by the presence

of many tight, highly refractile, hemispherical colonies in which individual cells cannot be distinguished. Human ES cell colonies are flatter, but should still have well-defined borders and contain many homogeneous, small, tightly packed cells. If starting cultures contain a majority of differentiated cells (spreadout, flattened colonies with ragged borders), then EB formation and lung differentiation efficiency will be suboptimal. It is impossible to eliminate spontaneously differentiating colonies entirely from ES cell cultures, but good culture technique will minimize this problem. Excellent instructions for the successful culture of human ES cells can be found at http://www.wicell.org. The inexperienced laboratory would be well advised to begin ES cell culture with murine cells, which are significantly easier to grow. It is also worth noting that EB cultures are particularly prone to contamination, as they are grown in petri dishes rather than filtered flasks. Take care not to leave a bridge of medium trapped between the base and lid of the dish, and handle the dishes carefully without tipping. To facilitate handling and provide an extra layer of protection, 10-cm dishes can be placed inside larger 14-cm dishes in the incubator. Basic Protocol 1 Apoptosis of the interior of the direct-contact coculture A disadvantage of using the whole EB for coculture is the fact that, after extended periods in culture, the cells within the interior begin to undergo apoptosis. This may be improved by placing the cultures on a rotating platform within the incubator to improve diffusion of gases and nutrients into the center of the construct. Failure of mesenchyme to fuse with EBs If the mesenchyme fails to fuse with the EB after 2 days of incubation, it may not be viable. This is likely to be due to excessive drying, which can occur during the dissection phase or initial overnight incubation. It can be avoided by ensuring that adequate volumes of medium are used to maintain the tissues throughout the procedure. Disaggregation of distal lung mesenchyme The authors’ experiments have also demonstrated that the disaggregation of the mesenchyme through trypsinization abrogates its ability to maintain SP-C expression. Epithelial rudiments of embryonic distal lung were

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cultured with trypsinized mesenchyme and intact mesenchyme pieces. Only the intact pieces were capable of maintaining SP-C mRNA expression after 5 days in culture. As a result, only experiments using whole mesenchyme pieces are performed. Epithelial contamination During the dissection process, it is possible to transfer epithelial pieces inadvertently along with the dissected mesenchyme. The features of the two tissues are fairly distinct under the microscope, but a few epithelial cells may go undetected. Therefore, it is necessary to be able to track the origins of the cells that are found to express the relevant markers after coculture. In the authors’ laboratory, this problem was overcome by using an ES cell line (E14Tg2a) that lacked hypoxanthine-guanine phosphoribosyltransferase (HGPRT) gene expression. In situ hybridization was performed using antisense HGPRT RNA probes, which did not localize to cells that expressed pro-SP-C/TTF-1 proteins, thus demonstrating their origins from ES cells. Ideally, a transgenic ES cell line that expresses a fluorescent reporter gene product (i.e., eGFP or DsRed) under the direction of the SP-C promoter should be used. The presence of the fluorescent protein would indicate the activation of SP-C gene transcription and the ES cell origin of the differentiated cells. The authors have generated such cells (see below) but not yet tested them in this coculture system. Basic Protocol 2 EBs fail to adhere during step 2 A proportion of EBs (up to 40%) will always remain in suspension, and these can be safely aspirated, as they do not appear to undergo significant levels of pulmonary epithelial differentiation. If EB attachment fails completely, it is possible that there is vibration in the incubator or that the plates have been disturbed too often. Under this protocol, EBs do not adhere firmly and can be easily dislodged within the first few days of plating onto gelatinized tissue culture plastic. However, the authors have observed that coating the tissue culture plastic with proteins that generate stronger EB adherence and spreading (e.g., Matrigel) tends to be detrimental to lung epithelial differentiation. Poor efficiency of lung epithelial differentiation In the authors’ laboratory, this method robustly generates around a 3% yield of lung epithelial progenitor cells in the viable cell

population. Where the yield has been lower, this has usually been a result of incorrect timing of the activin A treatment. Activin A must be added between days 2 to 3 of EB culture; later addition dramatically reduces lung differentiation efficiency. Reducing the concentration of activin A also reduces the end yield; the authors hypothesize that this is a consequence of mesoderm replacing endoderm specification in the early stages of EB differentiation. Lung marker proteins cannot be detected in differentiated ES cells This protocol generates early distal lung epithelial progenitor cells. At this stage of development, early lung markers such as SP-C and TTF-1 can be readily detected at the mRNA level but not at the protein level, and mature lung marker genes such as surfactant proteins A and B are yet to be induced. The authors routinely screen cultures by quantitative RT-PCR for phenotypic analysis and have generated a stably transfected murine ES cell line, containing the 4.8-kb murine SP-C promoter upstream of the eGFP gene (Rippon et al. 2006), in order to visualize and track lung epithelial progenitors in differentiating cultures. Basic Protocol 3 Time of changing to Dif ◦ hES medium It was noticed that, when EBs were transferred from nonadherent to adherent culture conditions, their adherence to the culture surface was affected by the timing of the switch from hEB medium to Dif◦ hES medium. When EBs were kept in the hEB medium for 2 days after their transfer into adherent culture conditions, more EBs attached to the surface than when medium was changed immediately after transfer of EBs into adherent culture conditions. Culture surfaces Three different culture surfaces were tested for attachment of differentiating hES cells: soda lime glass, tissue culture–treated Permanox (Nunc) and tissue culture–treated highgrade polystyrene (Orange Scientific). Among these tested surfaces, tissue culture–treated high-grade polystyrene was found to support attachment of EBs better than the others.

Anticipated Results Basic Protocol 1 In the authors’ laboratory, the efficiency at which pro-SP-C (pro-surfactant proteinC)/TTF-1–immunoreactive epithelial tubules (Fig. 1G.1.3) were obtained in the cocultures

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Figure 1G.1.3 Immunohistochemistry performed on sections of a direct contact coculture after 5 days of incubation. Antibodies to (A) TTF-1 and (B) pro–surfactant protein C were used on these serial sections. Original magnification, 400×.

Differentiation of Distal Lung Epithelium

(Basic Protocol 1) varied between experiments (by between 10% and 50%). Although the differentiation of ES cells into type II pneumocytes, based on the expression of these two markers, was demonstrated, it was necessary to exclude the possibility of contamination of the direct-contact cocultures by lung epithelium (Van Vranken et al., 2005). The efficiency of type II pneumocyte generation within EBs in the indirect cocultures (demonstrated by RT-PCR for SP-C mRNA expression) was variable (∼2- to 50-fold greater than EBs alone). There was some indication that

the inclusion of distal lung epithelium with the mesenchyme improved the outcome. Basic Protocol 2 Basic Protocol 2 yields highly mixed cultures in which the majority of cells are not lung epithelium. This optimized method reliably yields 2% to 4% differentiation efficiency as assessed by eGFP fluorescence in a cell line containing the eGFP gene under the control of the 4.8-kb murine SPC promoter (Fig. 1G.1.4). Endogenous SP-C and TTF-1 mRNA expression should be easily detected by

1G.1.18 Supplement 2

Current Protocols in Stem Cell Biology

Figure 1G.1.4 Fluorescent micrographs showing the position of eGFP-positive distal lung epithelial progenitors in adherent EBs at day 21 of differentiation, according to Basic Protocol 2. The murine ES cell line used in this experiment was stably transfected with a reporter cassette containing the eGFP gene under the control of the 4.8-kb murine surfactant protein C promoter. Original magnification, 200×.

quantitative RT-PCR in total RNA extracted from the whole mixed population. Under control conditions using serumcontaining medium, spontaneous differentiation of murine ES cells to TTF-1/SP-Cexpressing cells is very rarely detectable by qRT-PCR. Cultures allowed to differentiate spontaneously in EBDM alone are a useful negative control; additionally, non–activin A– treated controls can also be used to test the endoderm-induction step. Basic Protocol 3 This protocol results in a mixed population of differentiated cells, among which pulmonary epithelial cells could be identified by detection of SP-C mRNA using RT-PCR, immunostaining for surfactant proteins, A, B, C, and D, and electron microscopic detection of lamellar bodies. Figure 1G.1.5 shows a section of a sample with differentiated cells immunostained for surfactant protein A. The cells are mostly arranged in clusters. As surfactant protein C is expressed exclusively by

alveolar type II cells, the yield of these cells can be determined by calculation of the percentage of cells that express this protein, which is expected to be ∼2% of the population. According to the real-time RT-PCR studies performed in the authors’ laboratory, with the differentiating hES cells maintained in Dif◦ hES medium from 5 to 28 days, the SP-C mRNA expression level tends to increase. However, as the overall SP-C mRNA level is low, the large variation between samples precludes statistical analysis.

Time Considerations Basic Protocol 1 The coculture of EBs with murine embryonic distal lung epithelium takes up to 22 days to complete. An incubation of 8 to 10 days is required for the spontaneous differentiation of cells within EBs to endoderm. The EBs have been cocultured for up to 12 days, but this may be extended in optimal culture conditions. The actual process of dissecting the

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Figure 1G.1.5 A cluster of differentiated human ES cells immunostained for surfactant protein A (SP-A) by the ABC method and counterstained with hematoxylin. Brown stain represents SP-A and blue represents cell nuclei. Original magnification, 100×.

mesenchyme and manipulating the construct takes some time. Once proficient, it is possible to produce 12 to 15 direct-contact cocultures per day, dissecting tissue from about 10 embryonic lungs. Basic Protocol 2 The induction of the distal lung epithelial phenotype using soluble factors may take up to 31 days, with an additional 7 to 14 days required to produce the initial murine ES cell cultures, depending on the scale of the experiment. This is a three-stage differentiation protocol that comprises a 10-day suspension culture of embryoid bodies, followed by 11-day adherent culture of embryoid bodies and a 4- to 10-day selection of lung epithelial progenitors with lung-specific medium. The appearance of surfactant protein C–expressing lung epithelial progenitors should be observed between days 17 and 21 of differentiation; this therefore represents the earliest time at which the cells can be harvested, though the yield may be lower than in cultures allowed to differentiate for longer. The strength of Basic Protocol 2 is its ease of use. No specialized expertise and equipment, beyond those normally necessary to culture ES cells, are necessary, and the labor time involved at each step is minimal.

Differentiation of Distal Lung Epithelium

Basic Protocol 3 From the time the undifferentiated colonies are harvested for embryoid body formation, it will take up to 24 days for the experiment to be completed. A period of 7 days should be

allowed for hES cells to differentiate spontaneously in the embryoid bodies, an additional 12 days for continued differentiation in adherent culture conditions, and 5 days for differentiation in SAGM medium. The time needed for growth and propagation of undifferentiated hES cells should also be taken into account. When a frozen vial of hES cells is thawed, it might initially take up to 4 weeks before a subconfluent culture well can be obtained and the cells can be split for the first time. However, subsequent passages will grow faster, and they might reach subconfluence in less than a week.

Literature Cited Alcorn, J.L., Smith, M.E., Smith, J.F., Margraf, L.R., and Mendelson, C.R. 1997. Primary cell culture of human type II pneumonocytes: Maintenance of a differentiated phenotype and transfection with recombinant adenoviruses. Am. J. Respir. Cell. Mol. Biol. 17:672-682. Ali, N.N., Edgar, A.J., and Samdikuchaksaraei, A. 2002. Derivation of type II alveolar epithelial cells from murine embryonic stem cells. Tissue Eng. 8:541-549. Beverley, S. M. 2001. Enzymatic amplification of RNA by PCR (RT-PCR). Curr. Protoc. Mol. Biol. 56:15.5.1-15.5.6. Bogue, C.W., Jacobs, H.C., Dynia, D.W., Wilson, C.M., and Gross, I. 1996. Retinoic acid increases surfactant protein mRNA in fetal rat lung in culture. Am. J. Physiol. 271:L862-L868. D’Amour, K.A., Agulnick, A.D., Eliazer, S., Kelly, O.G., Kroon, E., and Baetge, E.E. 2005. Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23:1534-1541.

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D’Amour, K.A., Bang, A.G., Eliazer, S., Kelly, O.G., Agulnick, A.D., Smart, N.G., Moorman, M.A., Kroon, E., Carpenter, M.K., and Baetge, E.E. 2006. Production of pancreatic hormoneexpressing endocrine cells from human embryonic stem cells. Nat. Biotechnol. 24:13921401.

Nakano, T., Kodama, H., and Honjo, T. 1996. In vitro development of primitive and definitive erythrocytes from different precursors. Science 272:722-724.

Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., and 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.

Phelps, D.S., and Floros, J. 1991. Localization of pulmonary surfactant proteins using immunohistochemistry and tissue in situ hybridization. Exp. Lung Res. 17:985-995.

Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Edwards, L.A., Read, L.C., Nishio, S.J., Weir, A.J., Hull, W., Barry, S., Styne, D., Whitsett, J.A., Tarantal, A.F., and George-Nascimento, C. 1995. Comparison of the distinct effects of epidermal growth factor and betamethasone on the morphogenesis of the gas exchange region and differentiation of alveolar type II cells in lungs of fetal rhesus monkeys. J. Pharmacol. Exp. Ther. 274:1025-1032. Fair, J.H., Cairns, B.A., Lapaglia, M., Wang, J., Meyer, A.A., Kim, H., Hatada, S., Smithies, O., and Pevny, L. 2003. Induction of hepatic differentiation in embryonic stem cells by co-culture with embryonic cardiac mesoderm. Surgery 134:189-196. Kawasaki, H., Mizuseki, K., Nishikawa, S., Kaneko, S., Kuwana, Y., Nakanishi, S., Nishikawa, S.I., and Sasai, Y. 2000. Induction of midbrain dopaminergic neurons from ES cells by stromal cell-derived inducing activity. Neuron 28:31-40. Keller, G.M. 1995. In vitro differentiation of embryonic stem cells. Curr. Opin. Cell. Biol. 7:862869. Kubo, A., Shinozaki, K., Shannon, J.M., Kouskoff, V., Kennedy, M., Woo, S., Fehling, H.J., and Keller, G. 2004. Development of definitive endoderm from embryonic stem cells in culture. Development 131:1651-1662. Leahy, A., Xiong, J.W., Kuhnert, F., and Stuhlmann, H. 1999. Use of developmental marker genes to define temporal and spatial patterns of differentiation during embryoid body formation. J. Exp. Zool. 284:67-81. Lowe, L.A., Yamada, S., and Kuehn, M.R. 2001. Genetic dissection of nodal function in patterning the mouse embryo. Development 128:18311843. Madsen, J., Tornoe, I., Nielsen, O., Koch, C., Steinhilber, W., and Holmskov, U. 2003. Expression and localization of lung surfactant protein A in human tissues. Am. J. Respir. Cell. Mol. Biol. 29:591-597. Mateizel, I., De Temmerman, N., Ullmann, U., Cauffman, G., Sermon, K., Van de Velde, H., De Rycke, M., Degreef, E., Devroey, P., Liebaers, I., and Van Steirteghem, A. 2006. Derivation of human embryonic stem cell lines from embryos obtained after IVF and after PGD for monogenic disorders. Hum. Reprod. 21:503-511.

Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18.

Plopper, C.G., St. George, J.A., Read, L.C., Nishio, S.J., Weir, A.J., Edwards, L., Tarantal, A.F., Pinkerton, K.E., Merritt, T.A., Whitsett, J.A., et al. 1992. Acceleration of alveolar type II cell differentiation in fetal rhesus monkey lung by administration of EGF. Am. J. Physiol. 262:L313-L321. Pryhuber, G.S. 1998. Regulation and function of pulmonary surfactant protein B. Mol. Genet. Metab. 64:217-228. Rippon, H.J., Ali, N.N., Polak, J.M., and Bishop, A.E. 2004. Initial observations on the effect of medium composition on the differentiation of murine embryonic stem cells to alveolar type II cells. Cloning Stem Cells 6:49-56. Rippon, H.J., Polak, J.M., Qin, M., and Bishop, A.E. 2006. Derivation of distal lung epithelial progenitors from murine embryonic stem cells using a novel three-step differentiation protocol. Stem Cells 24:1389-1398. Samadikuchaksaraei, A., Cohen, S., Isaac, K., Rippon, H.J., Polak, J.M., Bielby, R.C., and Bishop, A.E. 2006. Derivation of distal airway epithelium from human embryonic stem cells. Tissue Eng. 12:867-875. Shannon, J.M. 1994. Induction of alveolar type II cell differentiation in fetal tracheal epithelium by grafted distal lung mesenchyme. Dev. Biol. 166:600-14. Strayer, M.S., Guttentag, S.H., and Ballard, P.L. 1998. Targeting type II and Clara cells for adenovirus-mediated gene transfer using the surfactant protein B promoter. Am. J. Respir. Cell. Mol. Biol. 18:1-11. Sui, Y., Clarke, T., and Khillan, J.S. 2003. Limb bud progenitor cells induce differentiation of pluripotent embryonic stem cells into chondrogenic lineage. Differentiation 71:578-585. Sun, Y., Li, H., Yang, H., Rao, M.S., and Zhan, M. 2006. Mechanisms controlling embryonic stem cell self-renewal and differentiation. Crit. Rev. Eukaryot. Gene Expr. 16:211-231. Tada, S., Era, T., Furusawa, C., Sakurai, H., Nishikawa, S., Kinoshita, M., Nakao, K., Chiba, T., and Nishikawa, S. 2005. Characterization of mesendoderm: A diverging point of the definitive endoderm and mesoderm in embryonic stem cell differentiation culture. Development 132:4363-4374. Van Vranken, B.E., Romanska, H.M., Polak, J.M., Rippon, H.J., Shannon, J.M., and Bishop, A.E. 2005. Coculture of embryonic stem cells with pulmonary mesenchyme: A microenvironment

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that promotes differentiation of pulmonary epithelium. Tissue Eng. 11:1177-1187. Voorhout, W.F., Weaver, T.E., Haagsman, H.P., Geuze, H.J., and Van Golde, L.M. 1993. Biosynthetic routing of pulmonary surfactant proteins in alveolar type II cells. Microsc. Res. Tech. 26:366-373. Watkins, S. 1989. Immunohistochemistry. Curr. Protoc. Mol. Biol. 7:14.6.1-14.6.13. Xu, X., McCormick-Shannon, K., Voelker, D.R., and Mason, R.J. 1998. KGF increases SP-A and SP-D mRNA levels and secretion in cultured rat alveolar type II cells. Am. J. Respir. Cell Mol. Biol. 18:168-178. Yasunaga, M., Tada, S., Torikai-Nishikawa, S., Nakano, Y., Okada, M., Jakt, L.M., Nishikawa, S., Chiba, T., Era, T., and Nishikawa, S. 2005. Induction and monitoring of definitive and visceral endoderm differentiation of mouse ES cells. Nat. Biotechnol. 23:1542-1550. Zeller, R. 1989. Fixation, embedding, and sectioning of tissues, embryos, and single cells. Curr. Protoc. Mol. Biol. 7:14.1.1-14.1.8.

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Pancreas Differentiation of Mouse ES Cells

UNIT 1G.2

Suzanne J. Micallef,1 Xueling Li,1 Andrew G. Elefanty,1 and Edouard G. Stanley1 1

Monash University, Victoria, Australia

ABSTRACT This unit describes the derivation of pancreatic cells from mouse embryonic stem cells (ESCs). Mouse ESCs are pluripotent immortal cells derived from the inner cell mass of pre-implantation blastocyst-stage embryos that possess the ability to differentiate into any cell type within the adult animal. In vitro, ESCs can be differentiated into a variety of cell types representing derivatives of the three embryonic germ layers, mesoderm, endoderm, and ectoderm. Successfully differentiating ES cells to pancreatic cells has the potential to provide an alternative to cadaver-derived cells for treatment of type I diabetes. This unit outlines a method for the differentiation of ESCs toward pancreatic endoderm in serum-free medium from embryoid bodies (EBs) formed in suspension or spin EBs. In addition there is a protocol for maintaining ESC. Curr. Protoc. Stem Cell C 2007 by John Wiley & Sons, Inc. Biol. 2:1G.2.1-1G.2.8.
Keywords: ES cells r differentiation r pancreatic endoderm

INTRODUCTION This unit describes the derivation of pancreatic cells from mouse embryonic stem cells (ESCs). Mouse ESCs are pluripotent immortal cells derived from the inner cell mass of pre-implantation blastocyst-stage embryos that possess the ability to differentiate into any cell type within the adult animal (Evans and Kaufman, 1981; Martin, 1981; Nagy et al., 1993). In vitro, ESCs can be differentiated into a variety of cell types representing derivatives of the three embryonic germ layers, mesoderm, endoderm, and ectoderm. This characteristic has led to the proposal that insulin-producing cells generated by the in vitro differentiation of ESCs might provide a limitless alternative to cadaveric-derived islets for the treatment of type I diabetes. However, given the large numbers of islets required to restore normoglycemia in people with type 1 diabetes (0.5 to 1 × 106 islet equivalents; Merani and Shapiro, 2006), it will be imperative to understand and optimize protocols for the pancreatic differentiation of ESCs. This unit outlines a method for the differentiation of ESCs toward pancreatic endoderm in serum-free medium (see Basic Protocol and Alternate Protocol). CAUTION: Retinoic acid is a teratogen. When working with this and other agents used in this protocol, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 8% CO2 incubator unless otherwise specified.

Embryonic and Extraembryonic Stem Cells Current Protocols in Stem Cell Biology 1G.2.1-1G.2.8 Published online July 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01g02s2 C 2007 John Wiley & Sons, Inc. Copyright


1G.2.1 Supplement 2

BASIC PROTOCOL

DIFFERENTIATION OF MOUSE ES CELLS TO PANCREATIC CELLS FROM EMBRYOID BODIES FORMED IN SUSPENSION This protocol describes the differentiation of ESCs to pancreatic endoderm using a multistep protocol in which cells are seeded at a defined density in low-attachment dishes.

Materials Mouse ESCs (Support Protocol) Mouse ES cell medium with LIF (see recipe) Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS) Trypsin-Versine chicken serum (TVCS; Invitrogen) Feeder depletion medium (see recipe) Chemically defined medium (CDM; see recipe) Recombinant human BMP4 (rhBMP4; R & D Systems) 10 mM all-trans retinoic acid (ATRA; Sigma-Aldrich) Trypan blue 1 M nicotinamide 15-ml conical centrifuge tube Refrigerated benchtop centrifuge (e.g., Sigma, 4K15; www.sigma-zentrifugen.de) 10-cm tissue culture dishes 6-cm low-adherent tissue culture dishes (Phoenix Biomedical). Adherent 96-well tissue culture plates 50-ml tubes Dissecting microscope Pulled, glass capillaries Additional reagents and equipment for cell counting (Phelan, 2006) Deplete feeder cells—day 0 1. On the day before a differentiation experiment is to begin and 1 hr before feeder depletion, exchange the medium on ∼80% confluent cultures of ESCs in 25-cm2 flasks with 5 ml fresh ES medium. One 25-cm2 flask should yield ∼5 × 106 ES cells and is sufficient for a standard differentiation.

2. Remove ES medium from the flask and wash with 5 ml of CMF-PBS. 3. Remove CMF-PBS and add 0.5 ml TVCS to the ESCs. Incubate for ∼2 min or until the ESCs easily detach from the surface of the flask with a gentle tap. 4. Add 5 ml of ES medium to inactivate the trypsin and pipet repeatedly to give a single-cell suspension. Transfer the cells to a 15-ml tube and centrifuge 5 min at 500 × g, 4◦ C. 5. Aspirate the supernatant and resuspend the cell pellet in 10 ml feeder-depletion medium. 6. Transfer the cell suspension to a 10-cm tissue culture dish and incubate for 40 min at 37◦ C in a tissue culture incubator. IMPORTANT NOTE: Do not disturb the dish during this time as the PMEFs will not adhere to the surface of the dish. At this time, the PMEFs will adhere to the tissue culture dish but the ESCs will not, allowing them to be collected in the supernatant. Pancreas Differentiation

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Initiate formation of EBs 7. Remove the dish from the incubator and carefully collect the supernatant (containing the ESCs) from the dish. The PMEFs will remain adhered to the surface of the dish.

8. Centrifuge the cell suspension for 5 min at 500 × g, 4◦ C and resuspend in 5 ml of CDM. 9. Perform a viable cell count using trypan blue (Phelan, 2006) and resuspend the ESCs at a density of 10,000 cells/ml in CDM. 10. Distribute 3 ml of the cell suspension per 6-cm low-adherent tissue culture dish. Add 2 ng/ml of recombinant human BMP4 to each dish (total of 6 ng/dish), mix by gently swirling each dish and place the dishes at 37◦ C in a tissue culture incubator. EBs are allowed to develop over the next 4 days. Approximately three dishes will yield 100 EBs of suitable size at day 5 of differentiation. At day 4 of differentiation EBs appear as a compact spherical mass of cells with a well-defined layer of endodermal cells around the border.

11. On day 4 collect EBs into 50-ml tubes (10 dishes/tube) and centrifuge 3 min at 500× g, 4◦ C. 12. Discard the supernatant and wash the EBs with 10 ml CMF-PBS. Centrifuge 3 min at 500 × g, 4◦ C, remove the supernatant carefully and resuspend in 15 ml fresh CDM. 13. Mix gently, and distribute 3 ml of the EB suspension per 6-cm low-adherent tissue culture dish.

Treat EBs with all-trans retinoic acid—day 4 14. Thaw ATRA at room temperature and add 3 µl/dish to make a final ATRA concentration of 10-5 M. Swirl dishes gently to mix the ATRA and place the dishes in a tissue culture incubator for a further 24 hr. Transfer EBs to adherent 96-well plates—day 5 15. On day 5, to prepare the 96-well plates, add 50 µl/well of the gelatin solution to each adherent 96-well plate required and incubate for 30 min at 37◦ C for 30 min. Aspirate the gelatin solution from each well and add 100 µl/well of CDM. 16. Collect the day 5 EBs into 50-ml tubes (10 dishes/tube) and centrifuge for 3 min at 500 × g, 4◦ C. 17. Discard the supernatant and wash the EBs with 10 ml CMF-PBS. Centrifuge for 3 min at 500 × g, 4◦ C, remove the supernatant carefully and resuspend in 9 ml fresh CDM. Transfer EBs to 3 × 6–cm tissue culture dishes. 18. Under a dissecting microscope, collect the EBs that are relatively uniform and large using a pulled glass capillary, transferring one EB at a time to each well of the prepared 96-well plates. Small EBs should be avoided as these do not yield pancreatic endoderm.

19. Place the 96-well plates in a tissue culture incubator, leaving the EBs to adhere and differentiate for a further 7 days. Pancreatic endoderm first appears at day 7 to 8 of differentiation, but requires time in culture to proliferate before the addition of nicotinamide. Embryonic and Extraembryonic Stem Cells

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Add nicotinamide to 96-well plate cultures—day 12 20. Aspirate the old CDM from the 96-well plates and replace with 100 µl fresh CDM containing 10 mM nicotinamide. Return the 96-well plates to the incubator and leave for 7 days. 21. Assess cells for markers of pancreatic differentiation. This is most easily achieved using genetically tagged ES cell lines, as described in Micallef et al., 2004 and 2007, where expression of Pdx1 and Insulin1 can be visualized by green and red fluorescent reporter genes, respectively. If using genetically unmodified ESCs, pancreatic differentiation should be assessed by assaying for expression of region specific markers (e.g., Pdx1 or Ptf1a), as well as markers representing differentiated endocrine (e.g., Insulin1, c-peptide, and pancreatic polypeptide) and exocrine (e.g., carboxypeptidase and amylase) cells. Expression of these markers can be analyzed by RT-PCR, immunohistochemistry, and transplantation studies. A more complete list of markers, antibodies, and PCR primers can be found in Micallef et al., 2004 and 2007. ALTERNATE PROTOCOL

DIFFERENTIATION OF ES CELLS TO PANCREATIC CELLS FROM SPIN EBS Due to the labor-intensive nature of “hand-picking” suitable EBs at day 5 of differentiation, a second method known as the “spin EB” method can be used to generate large numbers of EBs directly in 96-well plates (Ng et al., 2005b). This method also has the advantage that it generates higher numbers of pancreatic endodermal cells than the Basic Protocol.

Additional Materials (also see Basic Protocol) Gelatin solution Multichannel pipet Low-adherent, round-bottomed 96-well tissue culture plates Form spin EBs—day 0 1. Deplete ESCs of feeder cells as described in Basic Protocol (steps 1 to 9). 2. After the feeder depletion, resuspend the ESCs in CDM at a density of 2,500 cells/ml and transfer 100 µl of the cell suspension to each well of a 96-well plate to give a final cell density of 250 cells/well. The cell number has been optimized.

3. Place each 96-well plate in the plate holder of a benchtop centrifuge and aggregate the cells by centrifugation for 5 min at 500 × g, 4◦ C. This process forces the cells into a small cluster at the bottom of each well—a “spin EB.”

4. Incubate the spin EBs over the next 4 days in a tissue culture incubator.

Treat “spin EBs” with all-trans retinoic acid—day 4 5. On day 4, using a multichannel pipet, carefully remove half of the medium (50 µl) from each well of the 96-well plate, taking care not to disturb the EBs at the bottom of the well. 6. Thaw ATRA at room temperature and add 10 µl ATRA/5 ml CDM. Mix thoroughly, and add 50 µl of the ATRA/CDM mixture to each well (final concentration of ATRA is 10−5 M). Return the plates to the incubator for a further 24 hr.

Pancreas Differentiation

Transfer EBs to adherent 96-well plates 7. On day 5, prepare adherent 96-well plates by gelatinizing each well with 50 µl of 0.1% gelatin solution. Incubate for at least 30 min at 37◦ C before aspirating the gelatin solution just before the plates are required for use.

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8. Remove the spin EBs from the incubator. Aspirate as much of the medium as possible from the wells and then add 100 µl of fresh CDM to each well. 9. Using a multichannel pipet, transfer the EBs along with the 100 µl of fresh CDM to the adherent 96-well plates. Place the plates at 37◦ C in a tissue culture incubator. Culture the EBs for 7 days. At day 5 of differentiation, the ATRA is removed from the cultures and the EBs are transferred to adherent 96-well plates. As the spin EBs are initially cultured on lowadherent plates, they can be easily removed and transferred to gelatinized, adherent 96-well plates.

Add nicotinamide to 96-well plate cultures—day 12 10. Aspirate the old CDM from the 96-well plates and replace with 100 µl fresh CDM containing 10 mM nicotinamide. Place the 96-well plates at 37◦ C in a tissue culture incubator and leave EBs to differentiate for 7 days. Analyze for pancreatic markers, as described in step 21 of the Basic Protocol. MAINTENANCE OF EMBRYONIC STEM CELLS IN VITRO In order to preserve their stem cell phenotype, ESCs are maintained as adherent cultures in medium supplemented with leukemia inhibitory factor (LIF), often in the presence of mitotically inactivated primary mouse embryonic fibroblasts (PMEFs; Nagy et al., 2003). Upon removal of LIF and supporting PMEFs, ESCs can be induced to differentiate in suspension culture by the formation of embryoid bodies (EBs). This section describes the maintenance of undifferentiated ESCs in preparation for use in the differentiation procedures outlined above.

SUPPORT PROTOCOL

Materials Mitotically inactivated PMEFs PMEF medium (see recipe) Mouse ESC cultures grown on feeder cells in 25-cm2 tissue culture flasks Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS) Mouse ES cell medium with LIF (see recipe) Trypsin-Versine chicken serum (TVCS, Invitrogen) 25-cm2 gelatinized-tissue culture flask with vented cap (see recipe) 15-ml conical centrifuge tube Benchtop centrifuge (Sigma, 4K15; www.sigma-zentrifugen.de) 1. Seed mitotically inactivated PMEFs onto gelatin-coated 25-cm2 tissue culture flasks at a density of 4 × 104 cells/cm2 in 5 ml of PMEF medium. PMEFs should be prepared at least 24 hr prior to passaging ESCs.

2. When ESCs are ready to be passaged, remove the ES cell medium and wash with 5 ml of CMF-PBS. ESCs should be passaged every 2 to 3 days and the ES cell medium changed every day. ESCs should appear as uniform, refractile colonies with regular borders.

3. Remove CMF-PBS and add 0.5 ml TVCS to the ESCs. Incubate for ∼2 min at 37◦ C or until the ES cell colonies easily detach from the surface of the flask with a gentle tap. 4. Add 5 ml of ES medium to inactivate the trypsin and pipet repeatedly to give a single-cell suspension. Transfer the cells to a 15-ml tube and centrifuge 5 min at 500 × g, 4◦ C. Aspirate the supernatant and resuspend in 5 ml ES medium.

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5. Remove the PMEF medium from the flask of feeders and replace with 5 ml ES medium. Add the appropriate volume of ESCs to the flask and culture in an 8% CO2 tissue culture incubator. For general ES cell maintenance, ESCs should be passaged at a split ratio of approximately 1:10 to 1:15.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Chemically defined medium (CDM) 230 ml phenol-red free Iscove’s Modified Dulbecco’s Medium (IMDM; Invitrogen) 230 ml Ham’s F12 with Glutamax (Invitrogen) 33 ml 7.5% bovine serum albumin (BSA; Sigma, batch-tested) 5 ml 100× chemically defined lipids (1× final concentration) 5 ml 100× insulin, transferrin, selenium (ITS; 1× final concentration, Invitrogen) 500 µl LIF (1 U/µl final concentration; Chemicon) 19.5 µl monothioglycerol (MTG; 450 µm final concentration) Filter sterilize Store up to 1 week at 4◦ C Feeder-depletion medium 500 ml Iscove’s Modified Dulbecco’s Medium (IMDM; Invitrogen) 2.5 ml 10,000 U/ml penicillin/10,000 µg/ml streptomycin (from a 200× stock solution; Invitrogen) 50 ml fetal bovine serum (FBS) Filter sterilize Store for 1 month at 4◦ C Gelatin-coated tissue culture flasks and plates Prepare 0.1% (w/v) gelatin in PBS (Invitrogen) and autoclave. Store up to 1 month at 4◦ C. Incubate flasks or dishes with the gelatin solution for at least 30 min at 37◦ C, then remove gelatin and use immediately.

Mouse ES cell medium 411.5 ml DMEM (high glucose) 2.5 ml 10,000 U/ml penicillin/10,000 µg/ml streptomycin (from a 200× stock solution; Invitrogen) 5.0 ml 100× Minimal Essential Medium/Non-Essential Amino Acids (MEM/ NEAA; 10 mM final, Invitrogen) 5.0 ml 100× nucleosides (Sigma-Aldrich) 5.0 ml 100× L-glutamine (200 mM final concentration) 1 ml monothioglycerol (MTG; 50 mM final concentration; Sigma-Aldrich) 500 µl 106 U/ml LIF (Chemicon) 75 ml fetal bovine serum (FBS; batch selected, JRH Biosciences) Filter sterilize Store for 1 month at 4◦ C PMEF medium

Pancreas Differentiation

500 ml DMEM (high glucose) 2.5 ml 10,000 U/ml penicillin/10,000 µg/ml streptomycin (from a 200× stock solution; Invitrogen) 5.0 ml 100× L-glutamine (200 mM final concentration) continued

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50 ml fetal bovine serum (FBS; batch selected) Filter sterilize Store for 1 month at 4◦ C COMMENTARY Background Information ESCs are pluripotent cells that are derived from the inner cell mass of the blastocyst-stage embryo and have the potential to differentiate into any cell type within the adult, including cells of the pancreas. This method for the generation of pancreatic cells from ESCs utilizes a series of factors known to be important for pancreatic organogenesis. It has been demonstrated that BMP-4 alone is able to initiate a process resembling primitive streak formation in differentiating ESCs (Wiles and Johansson, 1999), exemplified by the expression of the primitive streak gene Mixl1, which is maximal at day 4 of differentiation (Ng et al., 2005a). Within the embryo, studies have shown that retinoic acid signaling during mid to late gastrulation is essential for later pancreatic development (Stafford and Prince, 2002; Stafford et al., 2004). Similarly, treatment of EBs with retinoic acid at day 4 of differentiation is also required to induce the optimal differentiation of ESCs to Pdx1-expressing pancreatic endoderm (Micallef et al., 2004). Finally, the addition of nicotinamide at day 12 of differentiation induces expression of insulin.

Critical Parameters and Troubleshooting The differentiation of ESCs in serum-free medium can be problematic, especially with respect to endoderm formation. In particular, the type and batch of BSA used in the CDM is critical. Testing of BSA is performed by assessing the ability of each batch to support retinoic acid–dependent induction of Pdx1 expression at day 12 using the Basic Protocol described in this unit. Criteria for batch testing FBS have been described previously (Barnett and Kontgen, 2001). Extensive testing of albumin types has shown that a 7.5% BSA solution from Sigma (no. A8412) is the best for inducing Pdx1+ endoderm; however, this still requires batch testing. Recombinant human albumin can also be used successfully (included in CDM at a concentration of 0.5%); however, it is currently not cost effective for routine use in standard experiments. In addition, the timing of the different steps throughout this protocol is paramount to its success. Variation in culturing techniques or

the different growth characteristics of ES cell lines may require some modification of the protocol. It is critical that ESCs are treated with ATRA when they are emerging from the in vitro equivalent of primitive streak formation. This can be established by determining the stage at which the EBs have maximal expression of the primitive streak genes Mixl1 or Brachyury. If EBs are treated with ATRA too early, induction of these genes is blocked, and differentiation will be diverted to an ectoderm cell fate (S.M., X.L., E.S., and A.E., unpub. observ.). Lastly, if cells are not being induced efficiently (see step 21 of the Basic Protocol) by BMP4, the concentration of BMP4 added to the cultures can be increased at day 0. When using the spin EB protocol (Alternate Protocol), the cell number added per well may require some optimization. Induction of pancreatic endoderm is ideal between 100 to 500 cells per well.

Anticipated Results By day 8 of differentiation, the emergence of some Pdx1+ endoderm can be detected. This population continues to proliferate and expand over the next 4 days, and by day 12 maximal Pdx1 expression can be detected. At this time, little or no insulin will be detected. Upon the addition of nicotinamide, insulin expression can be detected as soon as 2 days later; however, maximal insulin expression is not detected until day 20 of differentiation. In the absence of reporter ES cell lines such as the Pdx1GFP/w , ESCs in which green florescent protein (GFP) is expressed from the Pdx1 locus (Micallef et al., 2004), Pdx1+ and insulin+ cells can be detected by RT-PCR, flow cytometry, or immunohistochemistry.

Time Considerations The differentiation of mouse ESCs to pancreatic cells is a lengthy process that requires up to 20 days in culture. During this procedure, handling of the cells is required on days 0, 4, 5, and 12 of differentiation. This should be considered when setting up such experiments.

Literature Cited Barnett, L.D and Kontgen, F. 2001. Gene targeting in a centralized facility. Methods Mol. Biol. 158:65-82.

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Evans, M.J. and Kaufman, M.H. 1981. Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154-156.

Laboratory Manual (3rd edition). Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

Martin, G.R. 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. 78:76347638.

Ng, E.S., Azzola, L., Sourris, K., Robb, L., Stanley, E.G., and Elefanty, A.G. 2005a. The primitive streak gene Mixl1 is required for efficient haematopoiesis and BMP4induced ventral mesoderm patterning in differentiating ES cells. Development 132:873884.

Merani, S. and Shapiro, A.M. 2006. Current status of pancreatic islet transplantation. Clin Sci (Lond) 110:611-625. Micallef, S.J., Janes, M.E., Knezevic, K., Davis, R.P., Elefanty, A.G., and Stanley, E.G. 2004. Retinoic acid induces pdx1-positive endoderm in differentiating mouse embryonic stem cells. Diabetes 54:301-305.

Ng, E.S., Davis, R.P., Azzola, L., Stanley, E.G., and Elefanty, A.G. 2005b. Forced aggregation of defined numbers of human embryonic stem cells into embryoid bodies fosters robust, reproducible hematopoietic differentiation. Blood 106:1601-1603.

Micallef, S.J., Li, X., Janes, M.E., Jackson, S.A., Sutherland, R.M., Lew, A.M., Harrison, L.C., Elefanty, A.G., and Stanley, E.G. 2007. Endocrine cells develop within pancreatic bud-like structures derived from mouse ES cells differentiated in response to BMP4 and retinoic acid. Stem Cell Res. In press.

Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A3F.1-A.3F.18.

Nagy, A., Rossant, J., Nagy, R., AbramowNewerly, W., and Roder, J.C. 1993. Derivation of completely cell culture-derived mice from early-passage embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 90:8424-8428.

Stafford, D., Hornbruch, A., Mueller, P.R., and Prince, V.E. 2004. A conserved role for retinoid signaling in vertebrate pancreas development. Dev. Genes Evol. 214:432-441.

Nagy, A., Marina, G., Vintersten, K., and Behringer, R. 2003. Manipulating the Mouse Embryo: A

Stafford, D. and Prince, V.E. 2002. Retinoic acid signaling is required for a critical early step in zebrafish pancreatic development. Curr. Biol. 12:1215-1220.

Wiles, M.V. and Johansson, B.M. 1999. Embryonic stem cell development in a chemically defined medium. Exp. Cell Res. 247:241-248.

Pancreas Differentiation

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Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm

UNIT 1G.3

Alessandra Livigni,1 Santiago Nahuel Villegas,1 Ifigenia Oikonomopoulou,1 Afifah Rahman,1 Gillian M. Morrison,1, 2 and Joshua M. Brickman1 1

MRC Centre for Regenerative Medicine, Institute for Stem Cell Research, University of Edinburgh, Edinburgh, United Kingdom 2 Wellcome Trust Centre for Stem Cell Research, University of Cambridge, Cambridge, United Kingdom

ABSTRACT Anterior definitive endoderm (ADE) is both an important embryonic signaling center and a unique multipotent precursor of liver, pancreas, and other visceral organs. Here we describe a method for the differentiation of mouse embryonic stem (ES) cells to endoderm with pronounced anterior character. ADE-containing cultures can be produced in vitro by suspension (aggregation or embryoid body) culture and in a serum-free adherent monolayer culture. Purified ES cell–derived ADE cells appear committed to endodermal fates and can undergo further differentiation in vitro towards liver and pancreas with C 2009 by John enhanced efficiency. Curr. Protoc. Stem Cell Biol. 10:1G.3.1-1G.3.10.  Wiley & Sons, Inc. Keywords: embryonic stem cells r differentiation r endoderm r anterior r activin r FGF r Hex

INTRODUCTION Here we describe two protocols to differentiate mouse embryonic stem cells (mESCs) into definitive endoderm populations with a high degree of anterior character. These anterior definitive endoderm (ADE)–containing cultures can be produced by differentiation either in suspension culture (embryoid bodies) or as an adherent monolayer under serum-free conditions. From these differentiated cultures, it is possible to isolate cells that express high levels of ADE-specific markers and possess a gene expression profile that recapitulates definitive endoderm differentiation in vivo. mESC-derived ADE differentiates towards liver and pancreas with enhanced efficiency in vitro, while appearing committed to endodermal fates when challenged to differentiate towards other lineages both in vitro and in vivo. The first protocol (Basic Protocol, embryoid body differentiation) is a reproducible and efficient method to obtain ADE from embryoid bodies (EBs) and can be used for comparative studies alongside other EB protocols. The second protocol (Alternate Protocol, monolayer differentiation) is a defined protocol for ADE differentiation in an adherent monolayer format. This protocol employs the timed addition of specific cytokines, allows real-time observation of differentiation, and is particularly amenable to immunofluorescence analysis.

List of abbreviations ADE, anterior definitive endoderm; BMP4, bone morphogenetic protein 4; BSA, bovine serum albumin; DAPI, 4 ,6-diamidino-2-phenylindole; EBs, embryoid bodies; EGF, epidermal growth factor; FBS, fetal bovine serum, FGF4, fibroblast growth factor 4; mESCs, mouse embryonic stem cells; PBS, phosphate-buffered saline. Current Protocols in Stem Cell Biology 1G.3.1-1G.3.10 Published online July 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01g03s10 C 2009 John Wiley & Sons, Inc. Copyright 

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NOTE: All incubations should be performed in a humidified 37◦ C, 7% CO2 incubator. NOTE: Tissue culture procedures are performed in a laminar-flow hood using sterile equipment and tissue-culture grade solutions. All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: Before starting endodermal differentiation it is important to use cells that have had a chance to grow to a relatively high density (∼80% confluency). mESCs are routinely cultured as described in Li et al. (1995), Morrison and Brickman (2006), Zamparini et al. (2006), and Morrison et al. (2008). BASIC PROTOCOL

EMBRYOID BODY DIFFERENTIATION OF mESCs to ADE Embryoid body (EB) culture has been used routinely to assess differentiation towards a number of different lineages. Cells are grown in low-attachment vessels (bacteriological petri dishes) in GMEM medium in the absence of leukemia inhibitory factor (LIF). Under these conditions, ES cells form aggregates and begin the process of differentiation. These aggregates are cavitated, three-dimensional structures that contain all three germ layers, referred to as embryoid bodies (EBs). When cells are plated in suspension at low densities they form small aggregates (Kennedy et al., 1997). In the protocol described here, cells are plated at densities ∼100-fold lower than conventional EB protocols, allowing the formation of small, uniform aggregates (50 to 150 μm, Fig. 1G.3.1C) that ensure optimum cytokine exposure and promote anterior endoderm specification. These aggregates are allowed to grow for several days, and samples may be taken at various time points for analysis, e.g., by flow cytometry or RT-PCR. The protocol consists of two phases: LIF withdrawal in serum-containing medium, followed by further differentiation with activin and EGF under serum-free conditions.

Materials mESC culture medium (see recipe) without LIF mESC adherent culture in gelatin-coated 25-cm2 flask, ∼80% confluent (see Li et al., 1995; Morrison and Brickman, 2006; Zamparini et al., 2006; and Morrison et al., 2008, for culture technique) Calcium- and magnesium-free phosphate-buffered saline (CMF-PBS; Sigma, cat. no. D-8537) 0.025% trypsin (see recipe) N2B27 medium (Stem Cell Sciences; http://www.stemcellsciences.com/; thaw at 4◦ C and store up to 4 weeks at 4◦ C; if small amount is needed, refreeze in aliquots and store at –20◦ C until expiration date) Recombinant cytokine stock solutions (see recipe): Activin A EGF 75- or 150-cm2 sterile culture flask 15- and 50-ml sterile conical tubes Tabletop centrifuge Bacteriological petri dishes (Sterilin) Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3)

Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm

EBs day 0 1. Transfer an appropriate amount of mESC culture medium without LIF into an appropriately sized sterile flask or tube. Warm the medium aliquot in a 37◦ C water bath. Note that this flask will just be used to warm an aliquot of medium and not to culture cells.

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Figure 1G.3.1 Photomicrographs of cell aggregates in EB culture. (A) Bright-Þeld overview showing small aggregates generated after 2 days using the Basic Protocol. (B) Bright-Þeld overview showing growth of aggregates at day 4. (C) BrightÞeld overview showing regular ADE-enriched EBs generated after 7 days in culture by using the differentiation protocol (Basic Protocol). This protocol permits the generation of three-dimensional structures with spherical morphology, sized between 50 and 150 μm and highly enriched for ADE markers. (D) Fluorescence image from the same Þeld highlighting the induction of the ADE marker Hex in our Hex reporter ES cells. Red ßuorescence is produced by an enhanced dsRed targeted to the Hex locus, as evidenced by the expression of the Red Star protein. (E) Bright-Þeld image showing large aggregates or nonendodermal EBs. These aggregates indicate a failure of ADE differentiation. (F) Fluorescence image from the same Þeld, showing that these large EBs do not express Hex. Scale bars = 200 μm.

2. Aspirate the growth medium from an 80% confluent mESC adherent culture in a 25-cm2 flask, and rinse the cell layer twice, each time with 5 ml CMF-PBS. 3. Add 1 ml of 0.025% trypsin and incubate 1 min at 37◦ C, or until cells detach. Lightly tap the side of each flask to help detach cells from the surface.

4. Neutralize the trypsin by adding 9 ml of mESC culture medium without LIF and transfer the suspension to an appropriate conical tube. 5. Pipet gently up and down in order to form a single-cell suspension. 6. Centrifuge 2 min at 220 × g, room temperature, in a tabletop centrifuge.

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7. Remove the supernatant and resuspend the cells in 5 or 10 ml mESC medium without LIF. Be sure to obtain a single-cell suspension by pipetting gently several times.

8. Count the cells using a hemacytometer (UNIT 1C.3). 9. Seed the cells (5 × 103 /ml) in 10 ml of mESC medium without LIF onto a 10-cm bacteriological petri dish. 10. Gently agitate the dishes in order to get a uniform cell distribution.

EBs day 2 (48 hr after plating) 11. Collect EBs in appropriate conical tubes. 12. Centrifuge 1 min at 110 × g, room temperature. 13. Carefully remove the supernatant. Resuspend cells in 8 ml N2B27 medium containing 20 ng/ml Activin A and 20 ng/ml EGF, and transfer the EBs to a new 10-cm bacteriological Petri dish. 14. Gently agitate dishes to evenly distribute the EBs.

EBs day 4 (96 hr after plating) 15. Change the medium as on day 2 (repeat steps 11 to 14), resuspending the cells in 10 ml of N2B27 with 20 ng/ml cytokines (Activin A and EGF) per dish. 16. As EBs differentiate, collect them at various time points for analysis. Cells can be purified from day 6 or 7 cultures for further differentiation. ALTERNATE PROTOCOL

MONOLAYER DIFFERENTIATION OF mESCs TO ADE This protocol describes how to differentiate mESCs towards ADE as an adherent monolayer under defined, serum-free conditions. The lack of the complex three-dimensional interactions occurring in EBs, as well as the absence of serum, make this protocol ideal for the study of signaling pathways involved in both endoderm specification and patterning. It also allows real-time observation of differentiation. An example of monolayer differentiation is provided in Figure 1G.3.2.

Materials

Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm

N2B27 medium (Stem Cell Sciences; http://www.stemcellsciences.com/; thaw at 4◦ C and store up to 4 weeks at 4◦ C; if small amount is needed, refreeze in aliquots and store at –20◦ C until expiration date) 0.1% gelatin (see recipe) mESC adherent culture in gelatin-coated 25-cm2 flask, ∼80% confluent (see Li et al., 1995; Morrison and Brickman, 2006; Zamparini et al., 2006; and Morrison et al., 2008, for culture technique) Calcium- and magnesium-free phosphate-buffered saline (CMF-PBS; Sigma, cat. no. D-8537) 0.025% trypsin (see recipe) mESC culture medium (see recipe) without LIF Recombinant cytokine stock solutions (see recipe): Activin A BMP4 EGF FGF4 SFO3 medium (see recipe) 7.5% (75×) bovine serum albumin (fraction V; Invitrogen, cat no. 15260-037; store in aliquots at –20◦ C; once thawed, store at 4◦ C indefinitely)

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75- or 150-cm2 sterile culture flask 15- and 50-ml sterile conical tubes 6-well or 12-well tissue culture plates (Iwaki) Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3) Monolayer day 0 1. Transfer an appropriate amount of N2B27 medium into an appropriately sized sterile flask or tube. Warm the medium aliquot in the tissue-culture water bath at 37◦ C. Note that this flask will just be used to warm an aliquot of medium and not to culture cells.

2. Gelatinize an appropriate number of 6-well or 12-well tissue culture plates by adding enough 0.1% gelatin to cover the bottom of the well and leaving 10 min at room temperature. Aspirate the gelatin. 3. Aspirate the growth medium from an 80% confluent mESC adherent culture in a 25-cm2 flask and rinse the cell layer twice, each time with 5 ml CMF-PBS. 4. Add 1 ml of 0.025% trypsin and incubate 1 min at 37◦ C or until cells detach. Lightly tap the side of each flask to help detach cells from the surface. Avoid overtrypsinization, as this may result in poor cell survival in this protocol.

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Figure 1G.3.2 Photomicrographs of adherent monolayer culture. (A) Bright-Þeld image of monolayer culture on day 2 of differentiation. (B) Bright-Þeld image of monolayer culture on day 4 of differentiation. (C) Bright-Þeld overview showing regular morphology of ESCs that were differentiated using the Alternate Protocol after 7 days. ADE-enriched regions can be identiÞed by an abundance of the compact cells growing on top of the original monolayer. ADE is distributed throughout these areas of the dish. (D) DAPI nuclear staining. (E) Fluorescence image showing Hex-Red Star expression. Scale bars = 200 μm.

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5. Neutralize the trypsin by adding 9 ml of mESC culture medium containing serum (but without LIF), and transfer the suspension to an appropriate sized conical tube. 6. Pipet gently up and down in order to form a single-cell suspension. 7. Centrifuge 2 min at 220 × g in a tabletop centrifuge, at room temperature. 8. Resuspend the pellet in 10 ml of N2B27 medium (without cytokines) and count using a hemacytometer (UNIT 1C.3). Be sure to obtain a single-cell suspension by pipetting up and down gently several times.

9. Prepare an appropriate amount of N2B27 medium with cytokines: Activin A at 20 ng/ml, and BMP4 at 10 ng/ml. 10. Seed the cells at a density of 1 × 104 cells/ml in N2B27 medium with cytokines in gelatinized plates (see step 2) using 3 ml/well for 6-well plates or 1.5 ml for 12-well plates. Avoid swirling the plates, to avoid concentrating the cells at the center of the well. We recommend the use of 12-well plates for immunocytochemistry and 6-well plates for flow cytometry and RNA extraction.

Monolayer day 2 (48 hr after plating) 11. Transfer an aliquot of SFO3 medium into a sterile flask and warm it in the tissueculture water bath at 37◦ C. 12. Add to the SFO3: BSA (0.1% w/v; add from 75× stock), Activin A (20 ng/ml), EGF (20 ng/ml), and FGF4 (10 ng/ml). 13. Gently aspirate the N3B27 medium from the wells of the culture plates prepared in step 10. 14. Add SFO3 medium with cytokines to the wells (3 ml for 6-well plate, 1.5 ml for 12-well plate). Be gentle while using the vacuum and while pipetting, to avoid disturbing the cells attached to the edge of the well.

Monolayer day 4 (96 hr after plating) 15. Change the medium as on day 2 (repeat steps 10 to 14). Cytokines stored at 4◦ C can be reused within a week. No further medium changes are needed until day 7, when the anterior definitive endoderm population can be assessed either by molecular marker analysis or further differentiation.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Gelatin, 0.1% (w/v) Prepare a 1% stock solution of gelatin in tissue culture–grade water. Sterilize by autoclaving and divide into aliquots in sterile tubes. Dilute the stock to 0.1% in CMF-PBS (Sigma, cat. no. D-8537). Store up to 1 year at 4◦ C. Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm

mESC medium Glasgow MEM (Sigma, cat no. G5154) containing: 10% (v/v) FBS (see Troubleshooting for description of serum batch test) 0.1 mM nonessential amino acids

continued

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2 mM L-glutamine 1 mM sodium pyruvate 0.1 mM 2-mercaptoethanol Store up to 4 weeks at 4◦ C Just before use, for mESCs maintenance add: 1000 U/ml leukemia inhibitory factor (LIF; Chemicon, cat. no. ESG1107) Do not add LIF to the medium to be used in EB differentiation (Basic Protocol).

Recombinant cytokine stock solutions Resuspend lyophilized cytokines (all available from R&D Systems) in filtersterilized buffer (or HCl for BMP4) Stock concentrations: Activin A: 20 μg/ml in CMF-PBS (Sigma, cat. no. D-8537) with 0.1% (w/v) BSA EGF: 100 μg/ml in CMF-PBS with 0.1% (w/v) BSA BMP4: 10 μg/ml in 4 mM HCl FGF4: 100 μg/ml in CMF-PBS with 0.1% (w/v) BSA Store in small aliquots at –80◦ C according to manufacturer’s instructions. Once thawed, store up to 1 week at 4◦ C.

SFO3 medium Prepare the SFO3 medium (Iwai Chemicals; http://www.iwai-chem.co.jp/ company/index en.html) according to manufacturer’s instructions. Add 0.1 mM 2-mercaptoethanol. Filter sterilize. Store up to 4 weeks at 4◦ C

Trypsin, 0.025% (w/v) Mix 500 ml 1× CMF-PBS (Sigma, cat. no. D-8537) with 0.186 g disodium EDTA and filter sterilize. Add 5 ml of 2.5% trypsin (Invitrogen, cat no. 15090-046) and 5 ml chick serum (Sigma, cat no. C5405). Divide into aliquots in sterile tubes. Store up to 4 weeks at 4◦ C.

COMMENTARY Background Information Embryonic stem cells (ESCs) are karyotypically normal cell lines that can both self-renew indefinitely in vitro and generate all the lineages of both the fetus and the adult. When removed from conditions that promote selfrenewal, ESCs will generate progeny consisting of derivatives of the three embryonic germ layers: mesoderm, endoderm, and ectoderm (Smith, 2001; Nishikawa et al., 2007; Rossant, 2008). Recent advances in developmental biology have provided essential insights that have been used to recapitulate the differentiation of ESC towards specific lineages of interest. Embryoid body (EB) culture can be used to generate a number of different cell types, depending on the cell density and cytokine regime used. These in vitro–derived aggregates exhibit a three-dimensional structure that approximates early embryogenesis, and thus may reproduce a number of the interactions between different cell types and germ layers

and potentially allow for in vivo–like morphogenetic movements (Murry and Keller, 2008; ten Berge et al., 2008). To generate EBs, cells are grown in low-attachment dishes in medium without LIF. These conditions allow cells to form aggregates and commence differentiation. In mammalian development, endoderm induction occurs in two phases, the predominantly extra-embryonic visceral endoderm and the embryonic definitive endoderm (Beddington and Robertson, 1999; Lu et al., 2001). While recent data indicate that some visceral endoderm may contribute to the gut and visceral organs, these tissues are mostly constituted from the definitive endoderm (Kwon et al., 2008). As a result, considerable interest has been dedicated toward generating this lineage from ESCs. In embryos, the definitive endoderm forms first in the anterior region of the primitive streak, where it arises from an apparently homogenous group

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Differentiation of Embryonic Stem Cells into Anterior Definitive Endoderm

of mesendoderm cells. Cells in this region of the embryo express a uniform set of mesendoderm markers and have fates in both the mesoderm and endoderm lineages (Lewis and Tam, 2006). The first defined endoderm to migrate away from the primitive streak is the anterior definitive endoderm (ADE; Thomas et al., 1998). These cells will eventually constitute the ventral foregut and are thought to be multipotent precursors of liver and pancreas (Deutsch et al., 2001). The complexity of endoderm specification is highlighted by the fact that endoderm differentiation is extremely inefficient under standard EB conditions. However, reducing cell density and adding high levels of the Nodal related TGF-β activin promote endoderm formation (Kubo et al., 2004; Tada et al., 2005). While the cells of the mesendoderm region of the primitive streak contribute to all the visceral organs, they also contribute to the mesoderm and floor plate of the neural tube. To date, most strategies to obtain endoderm from ESCs have relied on using transgenic reporter cell lines that give a fluorescent readout of mesendodermal gene expression or panendodermal gene expression (visceral, definitive, anterior and posterior; Tada et al., 2005; Yasunaga et al., 2005; Gadue et al., 2006; Gouon-Evans et al., 2006). Differentiation was then optimized based on the expression of these reporters under a variety of conditions. As a result, these approaches did not optimize differentiation for the appearance of a positionally specified population, such as ADE. As previous protocols for ES cell differentiation to endoderm and endodermally derived cell types failed to progress through this positionally specified, physiological intermediate, we developed the protocols contained here for the generation of ADE. The ADE population can be isolated as a Hex- and Cxcr4positive population (Morrison et al., 2008). Hex is a homeobox transcription factor expressed in anterior definitive endoderm and in anterior visceral endoderm (Thomas et al., 1998; Brickman et al., 2000; Zamparini et al., 2006). Cxcr4 is a chemokine receptor expressed in mesoderm and endoderm but not in anterior visceral endoderm (McGrath et al., 1999; Yasunaga et al., 2005). The combination of these reporters was used to develop the protocols presented here. An example of Hexpositive ADE produced by both protocols is shown in Figures 1G.3.1 and 1G.3.2.

Critical Parameters and Troubleshooting Careful maintenance of mESC cultures is not only essential to preserve their pluripotency but also important for consistent and efficient endodermal differentiation. Cells should never be seeded to very low densities and never be allowed to overgrow past 90% confluency, in order to reduce spontaneous differentiation. Passage number is kept low (<50) by periodically thawing cells from liquid nitrogen stock. Plating or seeding differentiation experiments requires a single-cell suspension, and this is also critical for the efficiency of both the EB and the monolayer protocols. The seeding must be as gentle as possible. Cell culture technique during the plating process can affect the number of viable cells that commence differentiation, and this is a critical parameter for both protocols. However, as the monolayer occurs in a defined surface area, cell densities have a particularly pronounced effect on differentiation, and this can be compounded if the growth rate of different ESC lines varies during the protocol. Thus, poor results in a monolayer differentiation can often be rectified by changing the number of cells and the volume of medium in which they are plated. When experiencing problems, density can be raised to 2–3 × 104 cells/ml at seeding. We have also found that, in some instances, reducing the volume of medium by 33% can improve differentiation. Thus, for example, seeding 3 × 104 cells/ml in 1 ml for a 12-well plate is an alternative plating condition that we have used when troubleshooting. Seeding density is also important for EB formation. Small errors in the starting density may produce small EBs, but in high numbers. This can cause problems later on, as high densities of small EBs will aggregate to form large structures and inhibit ADE differentiation. This is best solved by adjusting the starting density, but it can also be solved by splitting the EBs collected at step 14 into twice as many plates. Another important factor to consider is the quality of the serum (fetal bovine serum; FBS). We usually batch test lots of FBS from different suppliers. We have found that mESC grown and/or differentiated in different sera can perform differently in EB differentiation. A simple way to screen sera is to examine whether the cells grown in a particular serum form uniform small aggregates (Fig. 1G.3.1C,D) or large sticky EBs (Fig. 1G.3.1E,F) upon

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differentiation. The small uniform aggregates are the ones that contain ADE at day 7, whereas the large sticky EBs do not. In a recent serum test, we found that about 1/3 of batch-tested sera were compatible with good endoderm differentiation. In our experience, the maximum percentage of cells with anterior endoderm markers is achieved at day 7 after plating. However, the factors mentioned above can affect the peak of induction. Therefore it is advisable to examine cultures at days 6 through 8 before attempting any large-scale differentiation.

Anticipated Results Following the EB protocol, you should observe small aggregates by day 2 (Fig. 1G.3.1A). These will then grow into modestly sized aggregates by day 4 (Fig. 1G.3.1B) and yield 200 to 300 EBs per plate by day 7. Up to 35% of this population expresses ADE markers by day 7, and ADE can be isolated as described by Morrison et al. (2008). ES cells differentiated using the monolayer differentiation grow as a single layer of adherent cells up to day 4 (Fig. 1G.3.2A,B). Late in the protocol, these cultures acquire a degree of three-dimensional character, with a layer of smaller cells growing on top of the original monolayer (see Fig. 1G.3.2C,D,E). ADE gene expression can be found in both cell layers, but the top layer appears to contain a higher proportion of Hex, Cer1–positive cells. After 7 days of differentiation using the monolayer protocol, the culture consists of ∼20% ADE. Monolayer culture can be monitored in real time by microscopy and at fixed time points by immunocytochemistry or by quantitative RT-PCR (Morrison et al., 2008). In a successful differentiation, ADE markers such as Hex and Cer1 are expressed from day 5 onwards. In particular, the coexpression of Hex and Cer1 with E-cadherin is reassuring. While Sox17 and Cxcr4 have been used as a marker for endoderm differentiation (see, for example, D’Amour et al., 2005, 2006; Yasunaga et al., 2005), these markers should not be used to judge whether ADE has been produced, and we have observed the expression of both markers as a result of a number of perturbations that completely abrogate anterior marker expression (data not shown). The progression of these cultures through particular stages of differentiation can be monitored by RT-PCR. For early primitive streak markers, we suggest Brachyury and Wnt3 (should peak at day 3), for anterior primitive streak, Goosecoid and

MixL1 (which should peak from days 4 to 6), and the ADE markers mentioned above. If the earlier primitive streak markers remain high in your cultures, it is a sign that your cultures either failed to progress or have produced large quantities of mesoderm (Morrison et al., 2008).

Time Considerations Obtaining an anterior definitive endoderm population should take 2 weeks. Once ESCs are thawed they will need about a week to recover. The differentiation protocols then requires 8 days, including the day when the cells are plated (day 0).

Acknowledgements We thank Rosa Portero Migueles and Maurice Canham for technical help and advice during the development of this unit. This work was supported by grants from the Biotechnology and Biological Sciences Research Council (BBSRC), Engineering and Physical Sciences Research Council (EPSRC), and Medical Research Council (MRC). J.M. Brickman is supported by a senior nonclinical fellowship from the MRC.

Literature Cited Beddington, R.S.P. and Robertson, E.J. 1999. Axis development and early asymmetry in mammals. Cell 96:195-209. Brickman, J.M., Jones, C.M., Clements, M., Smith, J.C., and Beddington, R.S.P. 2000. Hex is a transcriptional repressor that contributes to anterior identity and suppresses Spemann organiser function. Development 127:2303-2315. D’Amour, K.A., Agulnick, A.D., Eliazer, S., Kelly, O.G., Kroon, E., and Baetge, E.E. 2005. Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23:1534-1541. D’Amour, K.A., Bang, A.G., Eliazer, S., Kelly, O.G., Agulnick, A.D., Smart, N.G., Moorman, M.A., Kroon, E., Carpenter, M.K., and Baetge, E.E. 2006. Production of pancreatic hormoneexpressing endocrine cells from human embryonic stem cells. Nat. Biotechnol. 24:13921401. Deutsch, G., Jung, J., Zheng, M., Lora, J., and Zaret, K.S. 2001. A bipotential precursor population for pancreas and liver within the embryonic endoderm. Development 128:871-881. Gadue, P., Huber, T.L., Paddison, P.J., and Keller, G.M. 2006. Wnt and TGF-beta signaling are required for the induction of an in vitro model of primitive streak formation using embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A 103:16806-16811.

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Gouon-Evans, V., Boussemart, L., Gadue, P., Nierhoff, D., Koehler, C.I., Kubo, A., Shafritz, D.A., and Keller, G. 2006. BMP-4 is required for hepatic specification of mouse embryonic stem cell-derived definitive endoderm. Nat. Biotechnol. 24:1402-1411. Kennedy, M., Firpo, M., Choi, K., Wall, C., Robertson, S., Kabrun, N., and Keller, G. 1997. A common precursor for primitive erythropoiesis and definitive haematopoiesis. Nature 386:488-493. Kubo, A., Shinozaki, K., Shannon, J.M., Kouskoff, V., Kennedy, M., Woo, S., Fehling, H.J., and Keller, G. 2004. Development of definitive endoderm from embryonic stem cells in culture. Development 131:1651-1662. Kwon, G.S., Viotti, M., and Hadjantonakis, A.K. 2008. The endoderm of the mouse embryo arises by dynamic widespread intercalation of embryonic and extraembryonic lineages. Dev. Cell 15:509-520. Lewis, S.L. and Tam, P.P. 2006. Definitive endoderm of the mouse embryo: Formation, cell fates, and morphogenetic function. Dev. Dyn. 235:2315-2329. Li, M., Sendtner, M., and Smith, A. 1995. Essential function of LIF receptor in motor neurons. Nature 378:724-727. Lu, C.C., Brennan, J., and Robertson, E.J. 2001. From fertilization to gastrulation: Axis formation in the mouse embryo. Curr. Opin. Genet. Dev. 11:384-392. McGrath, K.E., Koniski, A.D., Maltby, K.M., McGann, J.K., and Palis, J. 1999. Embryonic expression and function of the chemokine SDF1 and its receptor, CXCR4. Dev. Biol. 213:442456. Morrison, G.M. and Brickman, J.M. 2006. Conserved roles for Oct4 homologues in maintaining multipotency during early vertebrate development. Development 133:2011-2022. Morrison, G.M., Oikonomopoulou, I., Migueles, R.P., Soneji, S., Livigni, A., Enver, T., and Brickman, J.M. 2008. Anterior definitive endo-

derm from ESCs reveals a role for FGF signaling. Cell Stem Cell 3:402-415. Murry, C.E. and Keller, G. 2008. Differentiation of embryonic stem cells to clinically relevant populations: Lessons from embryonic development. Cell 132:661-680. Nishikawa, S., Jakt, L.M., and Era, T. 2007. Embryonic stem-cell culture as a tool for developmental cell biology. Nat. Rev. 8:502-507. Rossant, J. 2008. Stem cells and early lineage development. Cell 132:527-531. Smith, A.G. 2001. Embryo-derived stem cells: Of mice and men. Annu. Rev. Cell Dev. Biol. 17:435-462. Tada, S., Era, T., Furusawa, C., Sakurai, H., Nishikawa, S., Kinoshita, M., Nakao, K., Chiba, T., and Nishikawa, S. 2005. Characterization of mesendoderm: A diverging point of the definitive endoderm and mesoderm in embryonic stem cell differentiation culture. Development 132:4363-4374. ten Berge, D., Brugmann, S.A., Helms, J.A., and Nusse, R. 2008. Wnt and FGF signals interact to coordinate growth with cell fate specification during limb development. Development 135:3247-3257. Thomas, P.Q., Brown, A., and Beddington, R.S.P. 1998. Hex: A homeobox gene revealing periimplantation asymmetry in the mouse embryo and an early transient marker of endothelial cell precursors. Development 125:85-94. Yasunaga, M., Tada, S., Torikai-Nishikawa, S., Nakano, Y., Okada, M., Jakt, L.M., Nishikawa, S., Chiba, T., Era, T., and Nishikawa, S. 2005. Induction and monitoring of definitive and visceral endoderm differentiation of mouse ES cells. Nat. Biotechnol. 23:1542-1550. Zamparini, A.L., Watts, T., Gardner, C.E., Tomlinson, S.R., Johnston, G.I., and Brickman, J.M. 2006. Hex acts with beta-catenin to regulate anteroposterior patterning via a Grouchorelated co-repressor and Nodal. Development 133:3709-3722.

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Differentiation of Mouse Embryonic Stem Cells to Spinal Motor Neurons

UNIT 1H.1

Hynek Wichterle1 and Mirza Peljto1 1

Departments of Pathology, Neurology and Neuroscience, Columbia University, New York, New York

ABSTRACT Controlled differentiation of embryonic stem (ES) cells into clinically relevant cell types is a fundamental goal of stem cell research. This unit describes one of the most efficient protocols for conversion of mouse ES cells into a defined type of nerve cells, the spinal motor neurons. ES cells are separated from feeder mouse embryonic fibroblasts and aggregated to form embryoid bodies (EBs). Two days after the withdrawal of growth factors, EBs reach a stage at which they are responsive to patterning signals and can be effectively induced with retinoic acid (RA) to differentiate into spinal nerve cells. Nascent neural cells become responsive to the ventralizing signal sonic hedgehog (Hh) that controls expression of ventral spinal progenitor markers and initiates the genetic program of motor neuron differentiation. Curr. Protoc. Stem Cell Biol. 5:1H.1.1-1H.1.9. C 2008 by John Wiley & Sons, Inc.
Keywords: controlled differentiation r mouse r embryonic stem cells r neural differentiation r spinal motor neurons r retinoic acid r sonic hedgehog

INTRODUCTION In this unit we describe a rapid and efficient protocol for differentiation of ES cells into spinal motor neurons. The protocol recapitulates the developmental and molecular processes that govern specification of motor neurons in the developing embryo (see Fig. 1H.1.1A; Wichterle et al., 2002). In the Basic Protocol we describe the differentiation procedure and in supporting protocols we discuss preparation of ES cells prior to the differentiation and methods for immunocytochemical characterization of differentiating motor neurons. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All centrifugations are performed in a swinging-bucket centrifuge at room temperature unless otherwise specified. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used throughout. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

DIFFERENTIATION OF ES CELLS INTO MOTOR NEURONS This protocol describes aggregation of ES cells into embryoid bodies (EBs) and directed differentiation of cells with retinoic acid (RA) and sonic hedgehog (Hh) to motor neurons. The progress and outcome of ES cell differentiation should be monitored by immunocytochemical analysis of developmentally regulated and lineage restricted markers (see Fig. 1H.1.1A).

Current Protocols in Stem Cell Biology 1H.1.1-1H.1.9 Published online May 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01h01s5 C 2008 John Wiley & Sons, Inc. Copyright


BASIC PROTOCOL

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1H.1.1 Supplement 5

Figure 1H.1.1 (A) Time line of ES cell differentiation. Temporal profile of expression of the key developmental markers is shown above the axis. Outline of the differentiation protocol is below the axis. (B) Typical shape of ES cell colonies (arrows) prior to trypsinization. Carry over fibroblast is marked with an asterisk. (C) Embryoid bodies at 2 days of differentiation. (D, E) Expression of motor neuron progenitor marker Olig2 (D) and post-mitotic motor neuron markers Hb9 and ISl1 (E) in immunostained sections of EBs. Abbreviations: ES, embryonic stem cells; PE, primitive ectoderm; NP, neural plate; pMN, progenitor motor neuron; MN, motor neuron; RA, retinoic acid; Shh, Sonic hedgehog protein; GDNF, glial cell line–derived neurotrophic factor; EBs, embryoid bodies.

Materials Dissociated ES cells suspended in ADFNK medium (see Support Protocol 1) ADFNK medium optimized for ES cell differentiation to HB9+ motor neurons (see recipe) 1 mM all-trans retinoic acid (RA; Sigma, cat. no. R2625) Sonic hedgehog (Shh) protein (R&D Systems); Purmorphamine (Calbiochem, cat. no. 540220); or Hh agonist HhAg1.3 (Curis, Inc.) Recombinant Rat GDNF (Glial cell line-derived neurotrophic factor; R&D Systems, 512-GF-050) 10-cm Nunc tissue culture dishes (Nunc, cat. no. 150679) Tissue culture microscope 15-ml tube Suspension culture dishes (10-cm; Corning, cat. no. 430591) 100-µm strainer (Falcon/Fisher Scientific, cat. no. 352360), optional 200-µl pipettor Large orifice pipet tips (Fisher Scientific, 21-197-2A) Additional reagents and equipment for fixation and immunocytochemistry (Support Protocol 2) Differentiation of mES Cells to Spinal Motor Neurons

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Differentiate ES cells 1. Plate 1 to 2 × 106 ES cells per 10-cm tissue culture dish in 10 ml of ADFNK (differentiation) medium. Evenly distribute cells by swirling the dish in a figure eight motion before placing the dish in the incubator. Although bacterial or suspension culture dishes can be used for differentiation, tissue culture treated dishes (such as Nunc Delta dishes) will improve the efficiency of differentiation by promoting selective adhesion of non-neural EBs during the first 2 days of differentiation.

2. On differentiation day 1 (Day 1) check the cultures for large number of EBs—small floating aggregates of ES cells that are visible under the microscope (a fraction of the aggregates will be adherent). Swirl the dish gently and transfer all floating EBs to 15-ml tube (discard the dish with adherent aggregates). Excessive adhesion of cells to tissue culture dishes indicates either that serum has been carried over to the differentiation medium or that ES cells are of poor quality and they prematurely differentiated along non-neural lineage.

3. Centrifuge EBs for 3 min at low speed (200 × g), room temperature. Aspirate supernatant, gently flick the tube to disperse EBs, add 10 ml of ADFNK medium and plate in a new 10-cm tissue culture dish. Swirl the dish to evenly distribute EBs and place back in the incubator.

Examine the cultures 4. On Day 2, examine the cultures: EBs should be of a fairly uniform size (50- to 100-µm in diameter) and most of them should be floating (see Fig. 1H.1.1C). Swirl the dish, collect EBs in a 15-ml tube and let them settle by gravity (∼15 min) or by gentle centrifugation (3 min at 200 × g, room temperature). Large aggregates of several EBs occasionally form during the first 2 days. If that happens, fit a 100-µm strainer in a 50-ml centrifuge tube and pipet the medium with EBs slowly in the center of the strainer. Transfer the flowthrough to a 15-ml tube and let the EBs settle as described above. Discard the strainer with large EB aggregates.

5. Aspirate medium and resuspend EBs in 400 µl of ADFNK medium. 6. Prepare four 10-cm culture dishes, each containing 10 ml of ADFNK medium supplemented with RA at 1 µM final concentration. 7. Using a 200-µl pipettor and a large-orifice yellow tip mix EBs by repeated pipetting and dispense 100 µl of EBs per 10-cm dish. Swirl dishes in a figure eight motion and return them to the incubator. Effective neuralization of EBs is achieved with RA concentrations ranging from 100 nM to 5 µM. RA is light sensitive and aliquots should be wrapped in aluminum foil and stored in the dark. To simplify the differentiation protocol, it is possible to supplement the medium with Hh on this day (see step 8). In particular, HhAg1.3 is relatively stable and will remain active until day 3 to 4 when it is required to ventralize EBs.

8. By day 3 differentiating ES cells reach the neural plate stage and can be patterned along the dorso-ventral axis. To induce motor neuron specification supplement dishes with Shh protein or Hh agonist (Shh protein at ∼200 ng/ml, Purmorphamine at 1 to 2 µM, or HhAg1.3 at 0.5- to 1 µM final concentration). Keep one dish supplemented only with RA as a negative control. The optimal concentration of Shh protein needs to be empirically determined, as individual batches vary. Purmorphamine is significantly less effective as compared to HhAg1.3 or Shh protein. On day 4 differentiating cells reach motor neuron progenitor stage marked by robust expression of Olig2 transcription factor (see Fig. 1H.1.1D).

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An aliquot of EBs can be collected at this stage for fixation and immunocytochemistry to determine the efficiency of motor neuron induction.

9. On day 5 check that the medium in the dishes has turned orange/yellow. Swirl dishes in a circular motion to collect EBs in the center of the dish (they are by now readily visible to the naked eye). Aspirate medium from the edges of the dish, taking care not to aspirate EBs, and add fresh ADFNK medium. Optionally, for better survival of motor neurons supplement the medium with GDNF to a final concentration of 5 ng/ml. 10. Day 6 to 7 is the peak of motor neuron differentiation. Collect EBs and fix for immunocytochemical analysis (Support Protocol 2) or for further culture and analysis of mature motor neurons (see Fig. 1H.1.1E). SUPPORT PROTOCOL 1

PREPARATION OF EMBRYONIC STEM CELLS FOR DIFFERENTIATION The efficiency of ES cell differentiation critically depends on the quality of ES cells. ES cells should be expanded under optimal growth conditions. Typical ES cell lines should be grown on monolayers of mitotically inactivated mouse embryonic fibroblasts in the presence of LIF and fetal bovine serum certified for ES culture (see UNIT 1C.3). The differentiation protocol assumes that expanded ES cells were frozen in aliquots each containing ∼2 million cells. This section describes recovery of frozen cells and preparation of ES cells for differentiation.

Materials Mouse ES cell line (for a better consistency it is recommended to start with a frozen aliquot of ES cells stored in liquid nitrogen) 70% ethanol ES medium (see recipe) ADFNK medium (see recipe) 0.05% (w/v) trypsin-EDTA (Invitrogen, cat. no. 05300-054) Gelatinized 25-cm2 culture flask (see recipe) 37◦ C water bath 15-ml tube Swinging-bucket tabletop centrifuge (Eppendorf, cat. no. 5702) Tissue culture microscope Gelatinized 75-cm2 culture flask, optional 10-cm tissue culture dish, optional Additional reagents and equipment for cell counting (Phelan, 2006) Recover ES cells 1. At a time point 2 days before differentiation, gelatinize 25-cm2 tissue culture flask, retrieve an aliquot of ES cells (∼2 million ES cells per aliquot) from the liquid nitrogen tank, and rapidly thaw the cells by submerging the vial in 37◦ C water bath until most of the ice melted. Wear goggles, as pressure will build up in the vial during thawing. Thawed ES cells should spend as little time as possible in DMSO-containing freezing medium.

2. Sterilize the vial with 70% ethanol and transfer into the laminar-flow hood. Gently mix the cells with 5 ml of warm ES medium in 15-ml tube and pellet in a clinical centrifuge (5 min at 200 × g, room temparature). Differentiation of mES Cells to Spinal Motor Neurons

While it is recommended to work quickly, freshly thawed ES cells are fragile and therefore avoid fast pipetting that generates shear stress.

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3. Aspirate supernatant, disperse ES cells by gently flicking the tube and resuspend in 5 ml of warm ES medium.

Count ES cells 4. Count ES cells using hemacytometer (Phelan, 2006) and plate 30,000 to 60,000 ES cells/cm2 in a gelatinized 25-cm2 flask in 6 ml of ES medium. Incubate the flask for 24 hr. Count only healthy looking ES cells that are small round and phase bright. Alternatively, use trypan blue to count viable, dye-excluding ES cells.

Remove fibroblasts 5. One day before differentiation remove the flask from the incubator, gently swirl the medium to dislodge cell debris and aspirate. Add 7 ml of warm ES medium, observe under the microscope, and return the flask into the incubator. Carryover embryonic fibroblasts should be attached and stretched on the substrate. ES cells should form small, compact islands of 2 to 8 cells with clearly delineated phase bright borders.

6. On the day of differentiation, examine the cultures. Replace ES medium with 6 ml of ADFNK medium 1 hr before trypsinization. ES cell colonies should appear mostly compact (see Fig. 1H.1.1B), covering 30% to 50% of the surface area.

7. Aspirate ADFNK medium and trypsinize ES cells with 3.5 ml of 0.05% trypsinEDTA (equilibrated to room temperature) in tissue culture incubator (2 to 5 min). Gently tap on the side of the flask and monitor dissociation under the microscope. 8. As soon as most colonies are floating, dilute trypsin by adding 5 ml of ADFNK medium, transfer cells to a 15-ml tube, and centrifuge 5 min at 400 × g, room temperature. 9. Aspirate medium and disperse colonies into single cells by firmly flicking the tube with a finger (∼10 times). Add 10 ml of ADFNK medium, mix by repeated pipetting, and count ES cells using a hemacytometer (Phelan, 2006). If excessive cell lysis occurred during trypsinization add DNaseI (10 µg/ml final concentration) to dissociated cells and mix by inverting the tube several times before centrifugation to prevent trapping dissociated cells in DNA. Five to ten million ES cells should be obtained from a single 25-cm2 flask.

10. Optional: If frozen vial of ES cells contained excessive number of fibroblasts it is recommended to reduce the amount of fibroblasts by selective adhesion to gelatincoated dishes. Resuspend trypsinized ES cells in 10 ml of ES medium, plate on gelatinized 75-cm2 flask or 10-cm tissue culture dish, and place in the incubator for 30 to 60 min. Fibroblasts should be firmly attached to the gelatinized surface while most ES cells should be still floating.

11. Gently swirl the dish and collect the medium containing floating ES cells. Centrifuge for 5 min at 400 × g, room temperature, and resuspend the cells in 10 ml of ADFNK medium. Repeat the centrifugation and resuspension in 10 ml of ADFNK to remove any remaining serum that would interfere with subsequent differentiation of ES cells. Count ES cells using a hemacytometer (Phelan, 2006). Embryonic and Extraembryonic Stem Cells

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SUPPORT PROTOCOL 2

FIXATION, CRYOSECTIONING, AND IMMUNOSTAINING During and at the end of the differentiation, the cells should be evaluated for the progress of differentiation. The simplest method of analysis is immunostaining of cryosections of EBs.

Materials Cultures of differentiating EBs (Basic Protocol) Phosphate-buffered saline (PBS; Cellgro, cat. no. 21-030-CV) 4% (w/v) paraformaldehyde (PFA; see recipe) 30% (w/v) sucrose in PBS Tissue-Tek OCT (Electron Microscopy Diatome, Fisher Scientific, 62550-12) Dry ice Blocking solution: 10% horse serum, 0.2% Triton X-100 in PBS Primary antibodies (see Table 1H.1.1) Secondary antibodies, fluorophore conjugated Aqua-Poly/Mount solution (Polysciences, cat. no. 18606) Wide-orifice pipet tip 1.5-ml microcentrifuge tubes Swinging-bucket tabletop centrifuge (Eppendorf, 5702) Embedding molds Superfrost Plus slides ImmunoPen (Calbiochem/EMD, cat. no. 402176) Humidified chamber Additional reagents and equipment for cutting sections using a cryostat (Watkins, 1989) and immunohistochemistry including slide processing (Hoffman, 2002) Fix cells 1. Collect EBs in the center of the dish by swirling the dish in a circular motion. Using a wide-orifice pipet tip, transfer EBs to a microcentrifuge tube. 2. Rinse EBs with PBS and fix for 30 to 90 min in fresh 4% PFA fixative on ice. Work in the fume hood as PFA is a carcinogen.

3. Wash EBs three times for 5 min each time with ice-cold PBS. 4. Cryoprotect EBs in 30% sucrose in PBS ∼60 min on ice. Wait until EBs equilibrate with sucrose and sink to the bottom of the tube. To prevent retention of EBs at the surface of the sucrose it is recommended to leave 0.5 ml of PBS in the tube and to layer 0.5 ml of sucrose at the bottom of the tube. EBs will stay at the interphase until they equilibrate with sucrose. Do not store EBs in sucrose for >12 hr as they will disintegrate. Table 1H.1.1 Antibodies Used for Screening Cells in Motor Neuron Differentiation

Differentiation of mES Cells to Spinal Motor Neurons

Antigen

Expression

Supplier

Catalog number

Recommended dilutions

Oct4

Expressed in ES cells

Abcam

ab19857

1:2000

Sox2

Expressed in ES cells- day 4

Chemicon

AB5603

1:2000

Pax6

Expressed day 3 – 4

DSHB

PAX6

1:30

Olig2

Expressed day 4 – 5

Chemicon

AB9610

1:20,000

Hb9

Expressed day 5 – 7

DSHB

81.5C10

1:100

Isl1

Expressed day 5 – 7

DSHB

39.4D5

1:100

Lhx3

Expressed day 5 – 7

DSHB

67.4E12

1:100

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5. Aspirate sucrose and PBS and add ∼1ml of Tissue-Tek OCT. Gently mix EBs with OCT (using a yellow tip) and spin them down in a swinging-bucket rotor (∼10 min at 500 × g, room temperature). 6. Transfer EBs into Tissue-Tek OCT-containing embedding mold and freeze on dry ice. To transfer EBs use large-orifice pipet tips. Eject EBs slowly to the bottom of the mold, making sure they do not disperse.

Cut sections 7. Using a cryostat, cut 10- to 15-µm thick sections (Watkins, 1989), collect them on Superfrost Plus slides and store the slides at −80◦ C until further analysis. Process slides 8. Process slides using standard immunocytochemical protocols (e.g., see Hoffman, 2002). Briefly, circle sections using an ImmunoPen and rehydrate in blocking solution. 9. Wash slides with PBS and incubate with 100 µl primary antibodies overnight at 4◦ C in a humidified chamber. 10. Wash slides three times, each time with PBS and incubate for 60 min with fluorophore-conjugated secondary antibodies. 11. Rinse slides three times, each time with PBS and add a coverslip using AquaPoly/Mount solution for fluorescent or confocal microscopy.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

ADFNK medium (differentiation medium) 250 ml Advanced DMEM/F12 (Invitrogen, cat. no. 12634-028) 250 ml Neurobasal medium (Invitrogen, cat. no. 21103-049) 5.6 ml Penicillin/streptomycin (from a 100 × stock solution purchased from Invitrogen, cat. no. 15140-155) 5.6 ml 200 mM L-Glutamine (Invitrogen, cat. no. 25030-081) 400 µl 1/100 diluted 2-mercaptoethanol in PBS (final 0.1 mM, Sigma) 56 ml Knockout serum replacement (Invitrogen, cat. no. 10828-028) Store in the dark in a cold room. Use within 3 weeks. ES medium 400 ml 75 ml

DMEM for ES cells (Chemicon, cat. no. SLM-220-B) ES cell tested fetal bovine serum (FBS; HyClone, cat. no. SH30070.03) or Knockout serum replacement (Invitrogen, cat. no. 10828-028). KSR results in better differentiation but ES cells do not grow that well. L-glutamine (Invitrogen, cat. no. 25030-081) 5 ml 5 ml 100× penicillin-streptomycin (Invitrogen, cat. no. 15140-122) 5 ml 100× non-essential amino acids (Chemicon, cat. no. TMS-001-C) 5 ml 100× Nucleosides (Chemicon, cat. no. ES-008-D,) 360 µl 1/100 diluted 2-mercaptoethanol in PBS (final 0.1 mM; Sigma, cat. no. M-7522) 50 µl LIF (final 1000 U/ml; Chemicon, cat. no. ESG1107) Store up to 1 week at 4◦ C

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Gelatinized flasks/plates Cover the bottoms of tissue culture flasks or plates with 0.1% (w/v) gelatin solution (Chemicon, ES-006-B). Incubate for >15 min at room temperature and aspirate gelatin. Gelatinized dishes can be stored up to 12 hr at room temperature.

Paraformaldehyde (100 ml), 4% Bring 90 ml of water to boil. In a fume hood add 4g of paraformaldehyde and 20 µl of 10 N NaOH. Stir for 30 min, add 10 ml of 10× CMF-PBS buffer (Chemicon, BSS-2010-B), pH 7.2, filter using a 0.22-µm filter and store at 4◦ C. Stable for up to 4 days.

COMMENTARY Background Information Neural differentiation The acquisition of motor neuron identity is controlled by two signaling molecules, retinoic acid (RA) and Hedgehog (Hh). RA has previously been shown to induce robust neural differentiation of ES cells grown as embryoid bodies (Bain et al., 1996). In addition, RA functions as a patterning signal that specifies the rostral spinal cord identity in vivo (Muhr et al., 1999). Once ES cells acquire spinal neural identity they become responsive to the ventralizing signal Hh that specifies among others motor neuron progenitor identity (Jessell, 2000). Differentiation of ES cells progresses in the same temporal sequence as differentiation of cells in the developing mouse embryo. Thus, efficient induction of spinal neural identity is achieved when embryoid bodies are treated with RA 2 days after the onset of differentiation, at a stage when cells acquire characteristics of primitive ectoderm. One day after the addition of RA, cells acquire early neural identity and can be patterned with Hh to induce expression of ventral neural markers and specify motor neuron progenitor identity (Wichterle et al., 2002). Differentiation of motor neuron progenitors into post-mitotic spinal motor neurons proceeds in a largely cellautonomous manner from day 5 to day 7 of differentiation.

Differentiation of mES Cells to Spinal Motor Neurons

Directed differentiation of human ES cells Protocols for directed differentiation of mouse ES cells were successfully extended towards directed differentiation of human ES (hES) cells into motor neurons (Li et al., 2005, 2008; Shin et al., 2005; Wilson and Stice, 2006; Lee et al., 2007). While the principals of differentiation driven by RA and Hh signals are similar for human and mouse ES cells, there are important differences that need to be considered. Mainly, generation of post-mitotic motor neurons from hES cells takes between

25 and 35 days in vitro instead of 6 days for mouse cells, highlighting the conservation of developmental time between normal embryonic development and in vitro differentiation. Synchronous differentiation of a population of hES cells over such a long period in culture is a challenge that likely contributes to the lower efficiency of human motor neuron production. An important step towards alleviating this hurdle has been recently made by synchronizing differentiation of hES cells at an intermediate neural plate–like stage characterized by neural rosette formation (Schulz et al., 2003; Wilson and Stice, 2006; Elkabetz et al., 2008).

Critical Parameters and Troubleshooting The quality of ES cells is one of the most critical parameters for efficient generation of motor neurons. Some ES cell lines are sensitive to fibroblast withdrawal and rapidly differentiate when plated on a gelatin-coated surface. Such lines should be grown on monolayers of feeder cells instead of gelatin-coated dishes. If an ES cell line is particularly prone to differentiation it is recommended to supplement ES medium with p38 kinase inhibitor SB203580 (1 µM, Calbiochem; Qi et al., 2004) or MEK inhibitor PD098059 (Calbiochem, 15 µM; Burdon et al., 1999), both of which prevent premature differentiation of ES cells. Both HhAg1.3 (Curis/Wyeth) and Shh protein (R&D Systems) are highly effective in ventralizing differentiating cells. In contrast, Purmorphamine (Calbiochem) is significantly less potent and should be used only if high efficiency of differentiation is not necessary. To monitor differentiation of ES cells into motor neurons it is possible to use HBG3 ES cell line carrying an Hb9-GFP transgene (Wichterle et al., 2002). Successful differentiation of ES cells into motor neurons can be monitored by live observation of EBs under the fluorescent microscope. Robust induction

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of GFP expression should be detectable from day 5 onwards.

Anticipated Results The differentiation protocol is suitable to generate large numbers of mature motor neurons. In a typical experiment 5 to 10 million ES cells are recovered after 2 days of growth on gelatin-coated dishes. One to two million ES cells are plated per 10-cm dish for differentiation. After splitting the dish 1:4 and induction with RA and HhAg1.3, the expected yield is ∼30 million cells on day 6 out of which ∼30% to 50% are motor neurons. Thus, if all ES cells were used for differentiation one can obtain ∼150 million cells or 50 to 75 million motor neurons per differentiation.

Time Considerations The entire motor neuron differentiation procedure for mouse ES cells takes 8 days. It is recommended to plate ES cells on Monday morning: ES cells are dissociated and transferred to differentiation medium on Wednesday and EBs are split in a 1:4 ratio and supplemented with RA or RA/Hh on Friday (Day 2). If EBs are supplemented with both RA and Hh on Friday the dishes can be kept undisturbed in an incubator over the weekend (otherwise supplement cultures with Hh on Saturday). Medium is changed again on Monday (Day 5) and mature motor neurons can be harvested for further experimentation or analysis on Tuesday or Wednesday.

Acknowledgements The authors would like to thank Dr. Thomas Jessell for his critical advice, encouragement, and support during the development of the described methods; and Curis Inc. for providing hedgehog agonist HhAg1.3. Annette Gaudino provided assistance with the preparation of manuscript. This work has been supported in part by Project A.L.S. Foundation.

Literature Cited Bain, G., Ray, W.J., Yao, M., and Gottlieb, D.I. 1996. Retinoic acid promotes neural and represses mesodermal gene expression in mouse embryonic stem cells in culture. Biochem. Biophys. Res. Commun. 223:691-694. Burdon, T., Stracey, C., Chambers, I., Nichols, J., and Smith, A. 1999. Suppression of SHP-2 and ERK signalling promotes self-renewal of mouse embryonic stem cells. Dev. Biol 210:30-43.

Elkabetz, Y., Panagiotakos, G., Al Shamy, G., Socci, N.D., Tabar, V., and Studer, L. 2008. Human ES cell-derived neural rosettes reveal a functionally distinct early neural stem cell stage. Genes Dev. 22:152-165. Hofman, F. 2002. Immuohistochemistry. Curr. Protoc. Immunol. 49:21.4.1-21.4.23. Jessell, T.M. 2000. Neuronal specification in the spinal cord: Inductive signals and transcriptional codes. Nat. Rev. Genet. 1:L20-29. Lee, H., Shamy, G.A., Elkabetz, Y., Schofield, C.M., Harrsion, N.L., Panagiotakos, G., Socci, N.D., Tabar, V., and Studer, L. 2007. Directed differentiation and transplantation of human embryonic stem cell-derived motoneurons. Stem Cells 25:1931-1939. Li, X.J., Du, Z.W., Zarnowska, E.D., Pankratz, M., Hansen, L.O., Pearce, R.A., and Zhang, S.C. 2005. Specification of motoneurons from human embryonic stem cells. Nat. Biotechnol. 23:215221. Li, X.J., Hu, B.Y., Jones, S.A., Zhang, Y.S., Lavaute, T., Du, Z.W., and Zhang, S.C. 2008. Directed Differentiation of Ventral Spinal Progenitors and Motor Neurons from Human Embryonic Stem Cells by Small Molecules. Stem Cells Jan 31, Epub ahead of print. Muhr, J., Graziano, E., Wilson, S., Jessell, T.M., and Edlund, T. 1999. Convergent inductive signals specify midbrain, hindbrain, and spinal cord identity in gastrula stage chick embryos. Neuron 23:689-702. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Qi, X., Li, T.G., Hao, J., Hu, J., Wang, J., Simmons, H., Miura, S., Mishina, Y., and Zhao, G.Q. 2004. BMP4 supports self-renewal of embryonic stem cells by inhibiting mitogen-activated protein kinase pathways. Proc. Natl. Acad. Sci. U.S.A 101:6027-6032. Schulz, T.C., Palmarini, G.M., Noggle, S.A., Weiler, D.A., Mitalipova, M.M., and Condie, B.G. 2003. Directed neuronal differentiation of human embryonic stem cells. BMC Neurosci. 4:27. Shin, S., Dalton, S., and Stice, S.L. 2005. Human motor neuron differentiation from human embryonic stem cells. Stem Cells Dev. 14:266-269. Watkins, S. 1989. Cryosectioning. Curr. Protoc. Mol. Biol. 7:14.2.1-14.2.8. Wichterle, H., Lieberam, I., Porter, J.A., and Jessell, T.M. 2002. Directed differentiation of embryonic stem cells into motor neurons. Cell 110:385-397. Wilson, P.G. and Stice, S.S. 2006. Development and differentiation of neural rosettes derived from human embryonic stem cells. Stem Cell Rev. 2:67-77. Embryonic and Extraembryonic Stem Cells

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Time-Lapse Imaging of Embryonic Neural Stem Cell Division in Drosophila by Two-Photon Microscopy

UNIT 1H.2

Elena Rebollo1 and Cayetano Gonzalez1,2 1 2

Cell Division Group, IRB-Barcelona, Barcelona, Spain Instituci´o Catalana de Recerca i Estudis Avanc¸ats, Barcelona, Spain

ABSTRACT This unit describes a protocol for live imaging of Drosophila embryonic neural stem cells using two-photon microscopy. Compared to traditional one-photon confocal imaging, this technique renders higher-resolution optical sections from deeper within the embryo. It is ideally suited to following embryonic neuroblasts located underneath the neuroepithelial cell layer for several rounds of cell division. Curr. Protoc. Stem Cell Biol. 13:1H.2.1C 2010 by John Wiley & Sons, Inc. 1H.2.9.  Keywords: two-photon microscopy r live imaging r Drosophila embryo r neuroblast

INTRODUCTION This unit provides a detailed description of how to prepare and image live Drosophila embryonic neural stem cells using two-photon microscopy. Drosophila embryos constitute an excellent system for live microscopy (Cavey and Lecuit, 2008) and have been used extensively to address the dynamics of different developmental processes. However, relatively little live-imaging data exist on the behavior of embryonic neuroblasts (NBs), the Drosophila neural stem cells, because as NBs delaminate from the embryonic neuroepithelium towards the interior of the embryo, they become inaccessible to standard microscopy techniques. Two-photon microscopy, which greatly improves resolution at deeper optical sections and causes little cell damage, can be used to circumvent this limitation. By fine-tuning the recording parameters, NBs can be followed through several consecutive divisions, providing an excellent tool to study different aspects of asymmetric cell division and embryonic neurogenesis. The unit begins with a Basic Protocol that describes the method, equipment, and parameters used to image embryonic NBs by two-photon microscopy. It is followed by a Support Protocol for collecting and preparing Drosophila embryos for live microscopy.

TIME-LAPSE IMAGING BY TWO-PHOTON MICROSCOPY This protocol describes the basic setup used to obtain a time-lapse series of images of dividing embryonic NBs using a two-photon microscope (see Video 1). Since we routinely use GFP-αTub84B and YFP-Asl (Rebollo et al., 2007) to visualize microtubules and centrioles, a description of how to excite and detect both GFP and YFP signals simultaneously is included. Microscope set-up conditions are given for a Leica TCS-SP5 multiphoton microscope (Leica Microsystems), but any multiphoton system that fits the minimum requirements described above (see Strategic Planning) can be utilized.

BASIC PROTOCOL

Materials Dechorionated embryos, mounted and ready for microscopy (see Support Protocol) Leica TCS-SP5 multiphoton LSM system (or equivalent), including:

Current Protocols in Stem Cell Biology 1H.2.1-1H.2.9 Published online June 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc01h02s13 C 2010 John Wiley & Sons, Inc. Copyright 

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General components: Inverted microscope (DMI 6000 CS) Laser point scanner system 63×/1.4 objective (oil immersion, HCX PL APO) Multiphoton components: Ti:sapphire laser Mai Tai (Spectra Physics) with wavelength range of 710 to 990 nm, pulse frequency of 80 MHz, giving maximum pulses of 1.5 W and 1 psec duration, and power control by EOM (electro-optical modulator) External detector (NDD, non-descanned detection): PMT 9624 (high sensitive meshless type, selected for Leica) Basic barrier filter SP720 (to stop infrared radiation and allow detection of whole GFP and YFP emission spectra) Leica LAS AF acquisition software (implemented for control of infrared laser) Software for visualizing and processing images (Imaris, Bitplane) and for assembling videos (Image J, http://rsb.info.nih.gov/ij/, or Adobe After Effects) Set up equipment 1. Switch on the confocal equipment and the pulse laser to warm them up and ensure that they are operative. It is best to set up the system before dechorionating any embryos, so that the preparation can be observed at the microscope immediately after it is mounted.

2. Make sure that the room temperature is maintained at a maximum of 23◦ C throughout the imaging session. Due to laser irradiation, the temperature at the sample will be higher than the room temperature. Embryos will die during recording if the room temperature reaches 26◦ C. For a more tight control of temperature, a temperature-controlled chamber around the sample may be used. The recommended chamber for the system described here is the Leica 11531834 (Cooling Thermostat, 230 V, control range –20◦ to 200◦ C, control tolerance ± 0.01◦ C).

3. Use the computer interface to select a wavelength for infrared excitation. To excite both GFP and YFP at the same time, 960 nm is efficient.

Focus microscope on embryo 4. Choose an embryo at the desired stage. To avoid photo-damage, bring the embryo into focus using transmitted light, which is also recommended to identify the developmental stage of the embryo. With practice, the 63× objective can be used for this purpose.

5. Once in focus, rotate the embryo to orient it for optimal image size. Avoid postacquisition rotations, which are time-consuming. Bear in mind that some software versions deliver digital images that are rotated by 90◦ with respect to the view through the microscope opticals (eyepieces).

6. Select the scanning format. For time-lapse acquisition, it is recommended to select small scanning formats, around 256 × 256 pixels. Larger formats produce more photo damage and slow down the acquisition rate. A 256 × 200 pixel format is recommended for recording embryonic NBs.

Time-lapse Imaging of Embryonic Neural Stem Cell Division

7. Select an intermediate laser power (∼50%), start scanning, and adjust laser gain and offset. It is not recommended to use more than 80% laser power, as this will produce too much heat within the specimen. The signal-to-noise ratio can be improved by using the “accumulate” function (which will add up the signal from three consecutive acquisitions)

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or by reducing the scanning speed to 200 Hz. The resulting signal can be digitally tuned by modifying the detector gain and offset. The under/overexposure option can be used to identify those areas of the image that are not properly exposed.

8. Select a region of interest and apply digital zoom to fit the image to the desired area. The ideal region to record Drosophila NBs is the ventral side of the embryo. Move the embryo until the ventral ectoderm (the more curved side) is in the center of the scanning field. Focus on the surface (neuroepithelial cell layer) and move in the z dimension until NBs appear in the field. Delaminating NBs can be recognized by their position (slightly off the epithelial layer), their shape (pear-like, with a protrusion that connects them to the epithelium), and later by the axis of division (perpendicular to the epithelial layer) and the asymmetry of division (rendering a large new NB and a small basal ganglion mother cell [GMC]). Since embryos contain several rows of delaminating NBs that are oriented from anterior to posterior, many NBs can be recorded at the same time. A 2.5 zoom factor will be enough to capture several NBs within the same field. Higher zoom factors will produce more damage during acquisition.

9. Re-adjust the image digitally using the under/overexposure function. 10. Select the upper and lower limits for the z stack to be acquired. Always acquire one extra section at both ends of the z stack. Make sure a maximum of 1 μm is set between sections. In this way, 30 μm can be easily covered in the z dimension, which allows for imaging NBs at different depths.

11. Select bidirectional scanning to speed up acquisition. 12. Activate the xyzt mode and adjust the interval frequency. The minimum time required for acquiring a stack of 30 sections will be 0.5 min with the specified settings. A 1-min interval is recommended to avoid continuous heating of the sample.

Acquire images 13. Start acquisition. During this process, external light sources may considerably increase image noise and must be avoided. Switch off the monitors and cover the microscope with a dark cloth.

14. To visualize the recorded four-dimensional (4-D) stacks and perform some basic image processing (selection, projection, filtering), use the LSM acquisition software or any other software able to handle 4-D stacks, such as Imaris (Bitplane). 15. To process videos from individual NBs, select the z sections corresponding to the cell of interest and project them using the maximum intensity projection function. The movie can be assembled using any software designed for such purposes, such as Image J or Adobe After Effects.

EMBRYO PREPARATION Optimal visualization of Drosophila embryos under the microscope requires removal of the chorion while leaving the viteline membrane intact, as the latter is transparent enough for fluorescence imaging and protects the embryo. Hand peeling is chosen instead of chemical dechorionation methods, because very few embryos are needed and chemical methods may affect viability. A brief summary is given below. Protocols for embryo collection and dechorionation—including recipes for apple plates and yeast paste and detailed descriptions of egg-laying cups—have been published elsewhere (Wieschaus and Nusslein-Volhard, 1988; Ashburner, 1989; Gonzalez and Glover, 2003; Kiehart et al., 2007; Rothwell and Sullivan, 2007).

SUPPORT PROTOCOL

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Also included is a description of how to mount the embryos for live microscopy. A standard protocol based on gluing the embryos to the cover glass and adding an oil layer on top to prevent desiccation is available (Cavey and Lecuit, 2008). However, to improve optical penetration, it is essential to minimize light scattering layers between the optics and the specimen. Therefore, it is better to mount the embryos directly onto the cover glass within a small drop of oil and cover them with an oxygen-permeable Teflon membrane that keeps them immobilized and slightly flattened (Kaltschmidt et al., 2000). Teflon membranes (YSI) are sold in packs of 100 as a component of oxygen monitors. Since they are oxygen-permeable and non-toxic, they can be used as a cheap alternative to Biofolie 25 (Heraeus), normally used for mounting live specimens for microscopy. Under these conditions, embryos can be kept alive and imaged for hours.

Materials Male and female Drosophila, 1 to 2 weeks old Apple plates: 3% (w/v) agar plates containing 4% (v/v) apple juice, freshly prepared and kept at 4◦ C Freshly made yeast paste Halocarbon oil (Sigma; halocarbon oil 700, cat. no. H8898 or Voltalef oil 10S) Plastic fly cages pierced with small holes to allow breathing Stereomicroscope Fine artist’s brush Double-sided cellophane tape (Scotch tape) Blunt forceps, e.g., standard No. 5 dissection forceps smoothed for this purpose 35-mm culture dish: Petri dish with a coverslip bottom (e.g., Fluoro Dish sterile culture dishes, World Precision Instruments, cat. no. FD#35) Teflon membranes (YSI Incorporated; cat. no. 5793) Vacuum grease (Dow Corning; cat. no. 976V) Collect embryos 1. Place well-fed females and males, ideally between 1 and 2 weeks old, inside a plastic fly cage. Any plastic beaker-like container that securely fits the diameter of the apple plates will work.

2. Seal the cage with an inverted apple plate supplemented with a drop of yeast paste at room temperature. Invert the cage so the plate is at the bottom, and keep in the dark at 25◦ C. Females lay better after being kept in such conditions for 1 or 2 days.

3. To start collecting, transfer flies to a clean cage with a new apple plate with fresh yeast. Change the plate every hour to make collections of nearly synchronized embryos. Label the plates with the collection time and allow the embryos to develop for 4.5 hr at 25◦ C.

Dechorionate embryos and prepare for microscopy The following steps are performed under a stereomicroscope. 4. Using a wet fine artist’s brush, take an embryo from the apple plate and place it on top of a piece of double-sided Scotch tape attached to a glass slide. Time-lapse Imaging of Embryonic Neural Stem Cell Division

5. Remove the chorion by gently rolling the embryo over the tape using the blunt tip of a pair of forceps. The dechorionated embryo will stick to the forceps.

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6. Transfer the embryo to a small drop of oil on the coverslip of a culture chamber. Any commercial brand can be used, provided the coverslip has the standard 0.17-mm thickness. To prevent desiccation, make sure that the embryo is fully covered by the oil.

7. Add two drops of vacuum grease at both sides of the oil drop using a fine pipet tip. 8. Cut a piece of Teflon membrane to a size that is just big enough to cover the oil and the vacuum grease drops. Place the membrane on top of the oil and press it down gently with the forceps, so that the embryo is immobilized and slightly flattened against the glass. The embryo is now ready for two-photon microscopy. When mounted as described, the embryo will naturally lay on the lateral surface, so that dorsal and ventral sides remain at the upper and lower edges, which is the ideal orientation for microscopy. It is advisable to process several embryos at once to ensure that at least one will be at the required developmental stage. NB delamination from the epithelium starts at stage 9 of embryogenesis, when embryos are ∼4.5 hr old. The preparation process should not take longer than 20 min. Thus, processed embryos should be between 3 hr 50 min and 4 hr 50 min old, right at or about to enter stage 9. This stage is easily recognized by landmarks that can be observed under a stereomicroscope, including the initial formation of the future stomodeal invagination (Wieschaus and Nusslein-Volhard, 1988).

COMMENTARY Background Information Neuroblast division Neuroblasts (NBs) are stem cell−like progenitors of the Drosophila nervous system. The first NBs delaminate from the polarized neuroectoderm early in development, at around stage 9 (Campos-Ortega and Hartenstein, 1997). Delamination is followed by asymmetric mitosis—with telophase figures aligned perpendicularly to the neuroectoderm—that results in a small differentiating ganglion mother cell (GMC) and a self-renewed NB. GMCs divide once more to produce cells that later differentiate into neurons or glia. Some larval NBs derive from quiescent embryonic NBs that re-enter the cell cycle. In both developmental stages, asymmetric division of NBs relies on the polarized localization of apical and basal protein complexes that are differentially segregated to the daughter cells by the controlled orientation of the mitotic spindle (Gonzalez, 2007; Knoblich, 2008). Pioneer time-lapse studies carried out in embryonic NBs revealed that the spindle is first assembled parallel to the neuroectoderm, and then rotates approximately 90◦ during metaphase to align with the apical/basal polarity axis (Kaltschmidt et al., 2000). More recent studies based on two-photon microscopy have shown that spindle rotation is limited to delaminating NBs. In later cell cycles, spin-

dles are assembled already aligned with the axis of cortical polarity (Rebollo et al., 2009). This predetermined spindle orientation mode was previously reported in larval NBs (Rebollo et al., 2007; Rusan and Peifer, 2007). Two-photon microscopy When optical sections are obtained deep within a sample using confocal microscopy, light scattering attenuates the fluorescence signal. Furthermore, scattering that occurs in regions away from the point of focus creates undesired fluorescence that will pass through the confocal pinhole to the detector, thereby increasing background. This deterioration in signal-to-noise levels is accentuated when using live fluorescence signals, which are generally less intense than fluorophores used in fixed samples. An alternative for imaging thick specimens is the two-photon system, in which excitation is caused by the simultaneous absorption of two photons with approximately twice the wavelength of the absorption peak of the fluorophore being used. In this manner, excitation is restricted to the focal plane, where the density of photons is high enough to allow two photons to simultaneously hit the fluorophore. An additional advantage of twophoton microscopy is that longer wavelengths reduce photo damage, thereby improving cell viability. Moreover, since no fluorescence is generated above or below the point of illumination, all detected signal can be used for

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two photon

one photon surface

A

B

C

D

E

Figure 1H.2.1

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imaging regardless of whether it has been scattered. More importantly, longer wavelengths suffer less scattering. Thus, the longer wavelengths allow deeper tissue penetration than can be achieved using the shorter excitation wavelengths used in standard confocal microscopy. Comparative views of a Drosophila embryo taken using two-photon versus onephoton imaging are shown in Figure 1H.2.1. Two-photon live imaging in Drosophila Live microscopy of Drosophila cells and tissues has been widely used to address many different questions related to cell behavior and division during development. It is relatively easy to create transgenic flies expressing different combinations of fusion proteins that render fluorescent chromosomes, kinetochores, centrosomes, microtubules, cell membranes, and so on in different colors and in the tissues of interest. This, together with the particular features of GFP derivatives (such as brightness, low toxicity, and the available palette of different excitation and emission spectra), makes these fluorescent markers invaluable for live microscopy in this system. Two-photon microscopy takes advantage of exactly these same properties. Drosophila embryos expressing fluorescent GFP reporters have been tracked using twophoton microscopy to address collective cell migration and movement during gastrulation (McMahon et al., 2008) and to analyze dendrite elimination in Drosophila dendritic arborizing sensory neurons during metamorphosis (Williams and Truman, 2005). The highintensity peaks of the pulse laser can also be used to ablate specific cells in the Drosophila embryo while preserving local integrity, which has made it possible to study the regulation of specific morphogenetic movements during development (Supatto et al., 2005). Two-photon excitation has become the imaging choice for highly light-scattering tissues,

and many more applications in Drosophila and other systems will be developed in the near future.

Critical Parameters and Troubleshooting Two-photon imaging can be directly implemented using a standard confocal laserscanning microscope. It requires a high peakpower pulse laser to ensure that the density of photons at the focal plane is sufficient to provoke a two-photon event, while the mean power levels remain moderate and do not damage the specimen. In addition, a special barrier filter that stops the infrared light reflected from the sample must be placed before the detector. Since two-photon excitation spectra are typically broader than the corresponding one-photon excitation spectra, it is possible to excite several fluorophores (e.g., GFP and YFP, or GFP and m-RFP) at a time with a single excitation wavelength. The emission spectra of both fluorophores may be detected as a single signal when such fluorophores are used to label cell structures that can be distinguished in other ways (for instance, by shape or localization); this requires that the barrier filter cube does not include any additional bandpass filter. In the protocol described here, microtubules and centrioles are readily distinguishable by structure, so GFP and YFP can be detected as a single signal. When fluorophores cannot be distinguished by the morphology of the labeled structures, spectral separation is required, using two external detectors with the appropriate bandpass filters. Using a unique excitation wavelength for several fluorophores will save time during the acquisition process and avoid cell damage due to extra laser power. Objectives with a high numerical aperture should be used to optimize photon concentration at the focal point. The infrared illumination point is moved along the sample by a

Figure 1H.2.1 (appears on previous page) Comparative view of two-photon versus one-photon imaging of the ventral neuroectoderm of a Drosophila embryo expressing GFP- and YFP-labeled microtubules and centrioles respectively. Two xyz stacks of 86 slices every 0.88 μm were acquired using two-photon and one-photon methods with a delay of 2 min between them. Selected z slices are shown at different depths from the embryo surface (12, 20, 42 and 70 μm). Very little difference is observed at the embryo surface (A) and at 12 μm (B), where dividing epithelial cells are observed (arrows in B). At 20 μm (C), differences in signal detection become very obvious. In delaminated NBs, the apical microtubule aster (arrowhead) and the basal cluster of daughter cells (asterisk) are perfectly distinguishable in the two-photon image, but are hardly visible in the corresponding one-photon image. At 42 μm (D), dividing NBs are still fully visible in the two-photon image (inset), but are undetectable in the corresponding one-photon image. Two-photon imaging can still render high-contrast images at 70 μm (E), where the epithelium is still observable. In contrast, there is a complete lack of signal with the one-photon system. Scale bar, 10 μm.

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computer-controlled scanner, which consists of mirrors located along the epiluminescence path. These mirrors move the laser beam in the xy plane along different z positions, allowing for three-dimensional reconstruction of the sample at each selected time point. A beam splitter is required to reflect the excitation infrared light to the objective and transmit the fluorescence emitted by the sample, which will be filtered again by the additional infrared barrier. Since two-photon excitation is restricted to the focal plane, detection pinholes are not needed. Nevertheless, the built-in detection pinhole can be used to reduce noise, although it will be at the expense of signal. Most microscopes have extra software tools to help adjust the image digitally. Due to the use of longer excitation wavelengths, resolution is always slightly lower in two-photon microscopy than in traditional confocal microscopy. The fluorescence emission intensity of a given fluorophore is also reduced in two-photon microscopy due to the pulsing activity of the laser. The loss of resolution can be partially compensated by reducing the aperture of the confocal pinhole, but at the expense of decreasing signal intensity. To compensate for this, it is possible to use lower scanning speeds or use functions that accumulate or average the signal and therefore reduce the noise. However, all of these manipulations will result in longer acquisition times and, ultimately, a loss of temporal resolution. Choosing a smaller scanning format will speed up the acquisition rate and may provide a compromise between spatial and temporal resolution. Increasing the repetition rate of the laser will also help improve signal intensity, with a limitation imposed by the viability of the specimen due to overheating. The use of objectives that are corrected for infrared light will definitely improve signal detection and resolution.

Anticipated Results

Time-lapse Imaging of Embryonic Neural Stem Cell Division

This protocol generates 4-D acquisitions of dividing embryonic Drosophila NBs that cannot be properly visualized with standard onephoton excitation techniques. The differences in penetration start to be evident at a 20-μm depth. The two-photon system allows one to follow cells throughout the entire z dimension of the embryo and to visualize several rows of NBs located at different depths. In the range of the specified magnification, ten to sixteen dividing NBs can be recorded at a time and over several consecutive divisions. The dura-

tion of acquisition and the final number of cell cycles followed will depend on the acquisition parameters. For long-term acquisitions, it is recommended to use longer time intervals (minimum 1 min).

Time Considerations The total time needed for embryo collection, development to the desired stage, and dechorionation and mounting for microscopy is approximately 6 hr. For a single embryo, a recording session of 2 hr will cover several rounds of division. If several consecutive embryo collections are done, two or three embryos can be followed in a single day. Processing will take much longer, even days, since the data volume produced is heavy and each stack will contain several NBs.

Acknowledgements We thank M´onica Rold´an for her valuable contribution to the establishment of the basic imaging conditions described in this protocol.

Literature Cited Ashburner, M. 1989. Drosophila: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Campos-Ortega, J.A. and Hartenstein, V. 1997. The Embryonic Development of Drosophila melanogaster. Springer, Berlin. Cavey, M. and Lecuit, T. 2008. Imaging cellular and molecular dynamics in live embryos using fluorescent proteins. Methods Mol. Biol. 420:219238. Gonzalez, C. 2007. Spindle orientation, asymmetric division and tumour suppression in Drosophila stem cells. Nat. Rev. Genet. 8:462-472. Gonzalez, C. and Glover, D.M. 2003. Techniques for studying mitosis in Drosophila. In The Cell Cycle: A Practical Approach (P. Fantes and R. Brooks, eds.), pp 143-175. IRL Press, Oxford. Kaltschmidt, J.A., Davidson, C.M., Brown, N.H., and Brand, A.H. 2000. Rotation and asymmetry of the mitotic spindle direct asymmetric cell division in the developing central nervous system. Nat. Cell Biol. 2:7-12. Kiehart, D.P., Crawford, J.M., and Montague, R.A. 2007. Collection, dechorionation, and preparation of Drosophila embryos for quantitative microinjection. Cold Spring Harb. Protoc. doi:10.1101/pdb.prot4717. Knoblich, J.A. 2008. Mechanisms of asymmetric stem cell division. Cell 132:583-597. McMahon, A., Supatto, W., Fraser, S.E., and Stathopoulos, A. 2008. Dynamic analysis of Drosophila gastrulation provides insights into collective cell migration. Science 322:15461550.

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Rebollo, E., Sampaio, P., Januschke, J., Llamazares, S., Varmark, H., and Gonzalez, C. 2007. Functionally unequal centrosomes drive spindle orientation in asymmetrically dividing Drosophila neural stem cells. Dev. Cell 12:467-474. Rebollo, E., Rold´an, M., and Gonzalez, C. 2009. Spindle alignment is achieved without rotation after the first cell cycle in Drosophila embryonic neuroblasts. Development 36:3393-3397. Rothwell, W.F. and Sullivan, W. 2007. Drosophila embryo dechorionation. Cold Spring Harb. Protoc. doi:10.1101/pdb.prot4826. Rusan, N.M. and Peifer, M. 2007. A role for a novel centrosome cycle in asymmetric cell division. J. Cell Biol. 177:13-20.

Supatto, W., Debarre, D., Moulia, B., Brouzes, E., Martin, J.L., Farge, E., and Beaurepaire, E. 2005. In vivo modulation of morphogenetic movements in Drosophila embryos with femtosecond laser pulses. Proc. Natl. Acad. Sci. U.S.A. 102:1047-1052. Wieschaus, E. and Nusslein-Volhard, C. 1988. Looking at embryos. In Drosophila: A Practical Approach. (D.B. Rogers, ed.), pp. 199-227. Oxford University Press, Oxford. Williams, D.W. and Truman, J.W. 2005. Cellular mechanisms of dendrite pruning in Drosophila: Insights from in vivo time-lapse of remodeling dendritic arborizing sensory neurons. Development 132:3631-3642.

Embryonic and Extraembryonic Stem Cells

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SECTION 2A Hematopoietic Stem Cells INTRODUCTION ematopoietic stem cells (HSCs) are defined by their ability to give rise to all lineages of the hematopoietic system, including red cells that transport oxygen and specialized cells that comprise the immune system. Experimental and clinical studies largely focus on the bone marrow compartment in adults to isolate HSCs, however, these rare cells can also be isolated from the peripheral circulation upon drug-induced mobilization from the bone marrow to the perpheral blood, or from neonatal umbilical cord blood, which contains transplantable HSCs. Due to the ease of harvest, many paradigms in stem cell biology have been founded on principles arising from the experimental and clinical study of HSCs and their function during in vivo transplantation. The importance of this section therefore extends beyonds HSC experimentation, since the editors and authors believe the value of these established methods will be of value in studying and newly defining stem cells from other tissue and sources.

H

Cord blood Cord blood is increasingly being harvested at birth and stored in banks around the world. Stocks are genetically diverse and therefore broaden the range of use so that these sources are applicable across human leukocyte antigen mismatches between donor and recipient. Although the number of HSCs in one cord blood sample seems currently insufficient to allow complete regeneration of the blood system for transplantation of an adult human, their capacity for proliferation and engraftment is reportedly greater than for HSCs derived from bone marrow. Cord blood HSCs also have a lower risk of infectious disease transmission. In addition to the number and quality of HSCs, successful transplantation depends on the presence of sufficient progenitors to fulfill short-term requirements for red cells, neutrophils, and macrophages before more primitive HSCs can contribute. Purification of HSCs and progenitors from cord blood facilitates their study and could in the long term enable their more efficient use in transplantation. In this section, the purification of HSCs from cord blood (UNITS 2A.2 & 2A.3) is described, as well as the isolation of mononuclear cells (UNIT 2A.1), which include progenitors as well as HSCs. Mick Bhatia and Roger Patient

Somatic Stem Cells Current Protocols in Stem Cell Biology 2A.0.1 Published online August 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a00s2 C 2007 John Wiley & Sons, Inc. Copyright


2A.0.1 Supplement 2

Isolation of Mononuclear Cells from Human Cord Blood by Ficoll-Paque Density Gradient

UNIT 2A.1

Taina Jaatinen1 and Jarmo Laine1 1

Finnish Red Cross Blood Service, Helsinki, Finland

ABSTRACT When preparing stem cell specimens from cord blood, pre-enrichment of mononuclear cells is highly recommended to improve the recovery of rare stem cells. Mononuclear cells are easily isolated by density gradient centrifugation. In Ficoll-Paque density gradient centrifugation, anticoagulant-treated and diluted cord blood is layered on the FicollPaque solution and centrifuged. During centrifugation, erythrocytes and granulocytes sediment to the bottom layer. Lower density lymphocytes, together with other slowly sedimenting cells such as platelets and monocytes, are retained at the interface between the plasma and Ficoll-Paque, where they can be collected and subjected to subsequent isolation of hematopoietic stem cells or to the culture of mesenchymal stem cells. Curr. C 2007 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 1:2A.1.1-2A.1.4.
Keywords: cord blood r fresh r cryopreserved r mononuclear cell r density gradient

When preparing stem cell specimens from cord blood, whole blood or mononuclear cell fractions can be used. However, pre-enrichment of mononuclear cells is highly recommended as it improves the recovery of rare stem cells. Mononuclear cells are easily isolated by density gradient centrifugation.

BASIC PROTOCOL

Enrichment of mononuclear cells by Ficoll-Paque density gradient centrifugation is used to improve the recovery of rare stem cells from whole blood. In Ficoll-Paque density gradient centrifugation, anticoagulant-treated and diluted cord blood is layered on the Ficoll-Paque solution and centrifuged. During centrifugation, erythrocytes and granulocytes sediment through the Ficoll-Paque to the bottom layer. Lower density lymphocytes, together with other slowly sedimenting cells such as platelets and monocytes, are retained at the interface between the plasma and Ficoll-Paque, where they can be collected and washed to remove platelets and residual Ficoll-Paque or plasma. The isolation of pure mononuclear cell fractions from cord blood, and subpopulations thereof, brings about a special challenge. In Ficoll-Paque density gradient centrifugation, all erythroid cells do not necessarily sediment to the bottom layer as they are expected to. Some erythroid cells may instead be retained in the interface of plasma and Ficoll-Paque. These cells are nucleated progenitors that are not easily depleted and may hamper the subsequent selection of stem cell populations. In addition, erythrocytes may form aggregates and adhere to lymphocytes, thus causing unusual sedimentation of lymphocytes in the bottom layer. Further, when handling cryopreserved cord blood, cell aggregation may occur due to cell damage during thawing. Means to reduce these problems are presented in this unit. Ficoll-Paque density gradient can be adapted to very small sample volumes. Thus, it is especially suitable for isolation of mononuclear cells from cord blood, as the sample size is often limited. Using density gradient centrifugation, mononuclear cells from a cord blood unit can be isolated within 2 hr. The mononuclear cell fractions prepared Current Protocols in Stem Cell Biology 2A.1.1-2A.1.4 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a01s1 C 2007 John Wiley & Sons, Inc. Copyright


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by density gradient centrifugation may be subjected to further selection procedures or mesenchymal stem cell cultures. Some cell loss may occur during density gradient centrifugation, but the method produces pure mononuclear cell fractions and removes nonviable cells effectively. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly.

Materials Anticoagulated cord blood (citrate dextrose, citrate phosphate dextrose, citrate, heparin, or EDTA) PBS/EDTA: Phosphate-buffered saline (PBS; see recipe)/2 mM disodium EDTA Ficoll-Paque Plus (Amersham Biosciences) or equivalent PBS/0.5% (w/v) bovine serum albumin/2 mM EDTA 500-ml Erlenmeyer flask or similar container 50-ml centrifuge tubes Centrifuge (preferably swinging-bucket rotor) Collect cord blood cells 1. Transfer anticoagulant-treated cord blood into a 500-ml Erlenmeyer flask or similar container and determine the blood volume. 2. Dilute blood 1:2 or preferably 1:4 with PBS/EDTA. Aggregation of erythrocytes is reduced by diluting the blood. When working with cryopreserved cord blood, it is recommended that the EDTA is replaced by anticoagulant citrate dextrose solution, formula A (0.6% ACD/A, Baxter Healthcare), which reduces aggregation more effectively. If the aggregation is substantial, the thawed cord blood cells may be pelleted by centrifuging for 10 min at 600 × g, 18◦ to 20◦ C, and then suspended in 200 µl of 1 mg/ml DNaseI (Sigma-Aldrich) to digest the DNA released from dead cells. Then the cells can be suspended carefully in 100 ml PBS supplemented with 0.6% ACD/A..

Separate mononuclear cells 3. Place 15 ml Ficoll-Paque solution into a 50-ml centrifuge tube. 4. Carefully layer 30 ml of the diluted blood on Ficoll-Paque solution. Use as many tubes as needed for the total sample volume. Do not mix blood and FicollPaque.

5. Centrifuge 40 min at 400 × g, 18◦ to 20◦ C, without brake. 6. Using a Pasteur pipet, collect the mononuclear cell fraction at the interface between plasma and Ficoll-Paque into a clean centrifuge tube. If there are many erythroid cells in the interface, treatment with 8% ammonium chloride or 3% diethylene glycol may be tested. Pellet cells by centrifuging 10 min at 700 × g, 18◦ to 20◦ C. Add 5 to 20 ml of lysis solution to the cell pellet, mix the suspension, and incubate 5 to 10 min at room temperature. Centrifuge 10 min at 700 × g, 18◦ to 20◦ C. Discard supernatant and proceed to step 7 (the depletion of erythrocytes may not be successful, as the nucleated erythroid progenitors are not easily lysed).

Wash the mononuclear fraction 7. Add 40 ml PBS/EDTA. Isolation of Mononuclear Cells from Human Cord Blood

8. Centrifuge 10 min at 300 × g, 18◦ to 20◦ C, with brake. 9. Discard the supernatant and repeat the wash with 40 ml PBS/EDTA. 10. Centrifuge 10 min at 300 × g, 18◦ to 20◦ C, with brake.

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Current Protocols in Stem Cell Biology

11. Discard the supernatant and suspend the mononuclear cells in 5 to 10 ml of PBS/0.5% bovine serum albumin/2 mM EDTA and proceed with cell counting. Continue with isolation of hematopoietic stem cells (UNIT 2A.2) or with the culture of mesenchymal stem cells (UNIT 2A.3). When working with cryopreserved cord blood, the aggregation may be so substantial that the cells need to be suspended in 200 µl of 1 mg/ml DNaseI (Sigma-Aldrich) to digest the DNA released from dead cells, thus preventing aggregation. Then PBS/0.5% bovine serum albumin/2 mM EDTA can be added in the appropriate volume (5 to 10 ml).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Phosphate-buffered saline (PBS) 0.23 g NaH2 PO4 (1.9 mM) 1.15 g Na2 HPO4 (8.1 mM) 9.00 g NaCl (154 mM) Add H2 O to 900 ml If needed, adjust to desired pH (usually 7.2 to 7.4) with 1 M NaOH or 1 M HCl Add H2 O to 1 liter Sterilize by filtering through 0.22-µm filter or by autoclaving Store indefinitely at 4◦ C Without adjustment, pH is normally ∼7.3. COMMENTARY Background Information The isolation of mononuclear cells from human blood using a low-viscosity erythrocyteaggregating agent was first described by Bøyum (Bøyum, 1964, 1968). There are many modifications for this method, but they all aim at easy, fast, and reproducible isolation of viable mononuclear cells. Several commercial solutions with proper density and viscosity are available for the density gradient centrifugation.

Critical Parameters and Troubleshooting The blood volume and tube diameter determine the height of the blood sample, which is critical to the successful isolation of a pure mononuclear cell fraction. Increasing the height of the blood sample augments erythrocyte contamination in the mononuclear cell fraction. A larger blood volume can be separated in a tube with larger diameter. Erythrocytes may also form aggregates and adhere to lymphocytes, thus leading to unusual sedimentation of lymphocytes in the bottom layer. This process can be reduced by diluting the blood. The more the whole blood is diluted, the better the separation of a pure mononu-

clear cell fraction. Also, a temperature of 18◦ to 20◦ C has been shown to give optimum results. All reagents should be adjusted to the temperature of 18◦ to 20◦ C.

Anticipated Results

Approximately a total of 1 to 3 × 108 mononuclear cells are obtained from one fresh cord blood unit. In fresh cord blood, the mean mononuclear cell concentration is 2.68 × 109 /liter, whereas it is 5.14 × 109 /liter in cryopreserved cord blood (Kekarainen et al., 2006). Cryopreserved cord blood is volumereduced, which explains the increase in cell concentration—number of cells is not increased. In the Finnish Cord Blood Bank, the final processed cord blood product contains 20 ml cord blood and 5 ml DMSO. The viability of recovered mononuclear cells is typically at least 95%.

Time Considerations The isolation of mononuclear cells from one cord blood unit is performed in 2 hr. The hands-on time is ∼1 hr.

Literature Cited Bøyum, A. 1964. Separation of white blood cells. Nature 204:793-794.

Hematopoietic Stem Cells

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Bøyum, A. 1968. Isolation of leucocytes from human blood. Further observations. Methylcellulose, dextran, and ficoll as erythrocyteaggregating agents. Scand. J. Clin. Lab. Invest. Suppl. 97:31-50.

Kekarainen, T., Mannelin, S., Laine, J., and Jaatinen, T. 2006. Optimization of immunomagnetic separation for cord blood-derived hematopoietic stem cells. B.M.C. Cell. Biol. 7:30.

Isolation of Mononuclear Cells from Human Cord Blood

2A.1.4 Supplement 1

Current Protocols in Stem Cell Biology

Isolation of Hematopoietic Stem Cells from Human Cord Blood

UNIT 2A.2

Taina Jaatinen1 and Jarmo Laine1 1

Finnish Red Cross Blood Service, Helsinki, Finland

ABSTRACT Enrichment of hematopoietic stem cells is based on the expression of certain surface antigens, such as CD34 and CD133, or on the lack of expression of lineage-specific antigens. Immunomagnetic positive selection of CD34+ or CD133+ cells is performed using paramagnetic microbeads conjugated to specific monoclonal antibodies (anti–human CD34 or anti–human CD133). In negative selection of lineage-negative (Lin− ) cells, the unwanted cells are labeled with antibodies against known markers for mature hematopoietic cells (CD2, CD3, CD14, CD16, CD19, CD24, CD56, CD66b, and glycophorin A) and retained in the column. Unlabeled cells pass through the column and are collected as the Lin− cell fraction. Immunomagnetic cell sorting system MACS is a fast and gentle method to enrich hematopoietic stem cells. Viable and highly pure cells can be separated to be used in various downstream applications, such as flow cytometry and cell culture. C 2007 by John Wiley & Sons, Inc. Curr. Protoc. Stem Cell Biol. 1:2A.2.1-2A.2.9.
Keywords: cord blood r fresh r cryopreserved r hematopoietic r CD34 r CD133 r Lin

INTRODUCTION Hematopoietic stem cells can be collected from peripheral blood, bone marrow, and cord blood mobilized by granulocyte colony-stimulating factor (G-CSF). Even though the cell content of cord blood is limited, it has a higher frequency of progenitor cells compared to peripheral blood or bone marrow (Broxmeyer et al., 1989; Grewal et al., 2003). Also, cord blood–derived CD34+ cells have been shown to proliferate more rapidly than their counterparts from bone marrow (Hao et al., 1995), and the cells have been shown to possess increased engraftment potential when compared to cells from peripheral blood or bone marrow (Vormoor et al., 1994; Hogan et al., 1997). Enrichment of hematopoietic stem cells is based on the expression of certain surface antigens, such as CD34 and CD133, or on the lack of expression of lineage-specific antigens. Immunomagnetic positive selection of CD34+ or CD133+ cells is performed using paramagnetic microbeads conjugated to specific monoclonal antibodies (anti-human CD34 or anti-human CD133). Labeled cells are enriched on a column placed in a magnetic field. Unlabeled cells pass through the column and the retained labeled cells can be eluted from the column after removal from the magnet. In negative selection or depletion, the unwanted cells are labeled with antibodies against known markers for mature hematopoietic cells and retained in the column. Unlabeled cells pass through the column and are collected as the lineage-negative (Lin− ) cell fraction. Mononuclear cells (UNIT 2A.1) are recommended as the starting material for immunomagnetic selection of hematopoietic stem cells. Aseptic cell processing is required if planning to perform cell culture experiments. Both fluorescence-activated cell sorting and immunomagnetic selection systems utilize antibodies against cell surface antigens. The immunomagnetic cell sorting system MACS is a fast and gentle method to enrich hematopoietic stem cells. Viable and functionally active cells can be separated to be used in various downstream applications, such as

Current Protocols in Stem Cell Biology 2A.2.1-2A.2.9 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a02s1 C 2007 John Wiley & Sons, Inc. Copyright


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flow cytometry and cell culture. Both labeled and unlabeled cells have good purity and recovery as well. The immunomagnetic selection can be made more effective using the AutoMACS system developed for high-speed automated cell sorting. The MACS column matrix provides a strong magnetic field, allowing the selection of cells carrying only few specific antigens on their surface, and the method is well suited for isolating rare stem cells. These protocols can be used to enrich CD34+/− , CD133+/− , or Lin−/+ cells with >90% purity from both fresh and cryopreserved cord blood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. BASIC PROTOCOL 1

ISOLATION OF CD34+ OR CD133+ CELLS FROM HUMAN CORD BLOOD Positive selection is used to enrich known hematopoietic stem cell fractions from human cord blood. The most commonly used surface markers for hematopoietic stem cell selection are CD34 and CD133. In immunomagnetic separation, CD34+ or CD133+ cells are labeled with magnetic microbeads attached to specific antibodies. The magnetically labeled cells are then purified and enriched in a magnetic field using MS or LS MACS columns, which are optimized for positive selection of cells. Both CD34+ and CD133+ cells have been used in stem cell transplantation and the number of CD34+ cells is used to depict the stem cell content of cord blood units in cord blood banking (Aroviita et al., 2005). A flow chart of the protocol steps is presented in Figure 2A.2.1.

Materials Mononuclear cells from human cord blood (UNIT 2A.1) Labeling buffer (see recipe), degassed Direct CD34 Progenitor Cell Isolation Kit (no. 130-046-702, Miltenyi Biotec) or CD133 Cell Isolation Kit (no. 130-050-801, Miltenyi Biotec) containing: FcR blocking reagent MicroBeads 10-ml centrifuge tubes Centrifuge (preferably a swinging-bucket rotor) MACS columns (MS, no. 130-042-201 or LS, no. 130-042-401; Miltenyi Biotech) MACS separator (MiniMACS, no. 130-042-102 or MidiMACS, no 130-042-302; Miltenyi Biotec) Additional reagents and equipment for preparing mononuclear cell suspension (UNIT 2A.1) Label the cells 1. Prepare mononuclear cell suspension (UNIT 2A.1) in a 50-ml centrifuge tubes. 2. Add 300 µl of labeling buffer per 108 mononuclear cells (for fewer cells use 300 µl of labeling buffer). Use degassed buffer only, as the gas bubbles may lead to clogging of the column and decrease the efficacy of immunomagnetic separation. EDTA in the labeling buffer may be replaced by other anticoagulants, such as 0.6% citrate dextrose or citrate phosphate dextrose. When working with higher cell numbers, scale up the labeling buffer and reagent volumes (e.g., 300 µl of labeling buffer per 108 mononuclear cells or 600 µl of labeling buffer per 2 × 108 mononuclear cells). Isolation of Hematopoietic Stem Cells from Human Cord Blood

2A.2.2 Supplement 1

3. Add 100 µl of FcR blocking reagent and 100 µl of MicroBeads per 108 mononuclear cells. Use CD34 or CD133 microbeads to isolate CD34+ or CD133+ cells, respectively. Use cold reagents and solutions to avoid capping of antibodies. Blocking reagent (human IgG) is added to inhibit unspecific or Fc-receptor-mediated binding of the MicroBeads. Current Protocols in Stem Cell Biology

Figure 2A.2.1 Protocol 1).

Flow chart of the protocol steps to isolate CD34+/− or CD133+/− cells (Basic

Current Protocols in Stem Cell Biology

Hematopoietic Stem Cells

2A.2.3 Supplement 1

4. Mix well and incubate at 4◦ to 12◦ C for 30 min. 5. Add 10 ml of labeling buffer. 6. Centrifuge for 10 min at 300 × g, 18◦ to 20◦ C. Discard supernatant from the wash. 7. Resuspend cells in 500 µl of labeling buffer per 108 mononuclear cells.

Perform magnetic separation 8. Place the appropriate column in the magnetic field of the MACS separator. Choose the column type according to the number of mononuclear cells; MS column for <2 × 108 cells and LS column for 2 × 108 to 2 × 109 cells.

9. Rinse the column with labeling buffer: 500 µl for MS column and 3 ml for LS column. 10. Apply labeled cell suspension (from step 7) to the column. If cell aggregates are formed, the cell suspension may be filtered using 30-µm nylon mesh or preseparation filters (no. 130-041-407, Miltenyi Biotech) before applying the cell suspension to the column.

11. Allow unlabeled cells to pass through the column and collect the flowthrough as the CD34− or CD133− cell fraction. The CD34− or CD133− cells are often collected for control purposes.

12. Wash the column with labeling buffer, four times with 500 µl for MS column and four times with 3 ml for LS column. 13. Remove the column from the magnetic field and place on a new centrifuge tube. 14. Elute and collect the CD34+ or CD133+ cells with labeling buffer: two times with 1 ml for MS column and two times with 5 ml for LS column. Use the plunger supplied with the column on the second elution.

Label cells 15. Centrifuge the eluted cells for 5 min at 300 × g, 18◦ to 20◦ C. 16. Discard the supernatant and resuspend the cells in 300 µl of labeling buffer. 17. Add 25 µl of FcR blocking reagent and 25 µl CD34 or CD133 MicroBeads. 18. Mix well and incubate at 4◦ to 12◦ C for 15 min. 19. Add 4 ml of labeling buffer. 20. Centrifuge for 5 min at 300 × g, 18◦ to 20◦ C. Discard the supernatant from the wash. 21. Resuspend cells in 500 µl of labeling buffer.

Perform magnetic separation step 22. Place MS column in the magnetic field of the MACS separator. 23. Rinse the column with 500 µl of labeling buffer. 24. Apply labeled cell suspension to the column. 25. Wash the column four times with 500 µl of labeling buffer. Isolation of Hematopoietic Stem Cells from Human Cord Blood

26. Remove the column from the magnetic field and place on a new centrifuge tube. 27. Elute and collect the CD34+ or CD133+ cells two times with 500 µl of labeling buffer. Use the plunger supplied with the column on the second elution.

2A.2.4

The cell suspension containing the CD34+ and CD133+ cells can be used in any laboratory assays or it can be frozen for later use.

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Current Protocols in Stem Cell Biology

ISOLATION OF LIN− CELLS FROM HUMAN CORD BLOOD Hematopoietic stem cells may be enriched based on the lack of lineage-specific antigens on their cell surface. To enrich progenitor cells, mononuclear cells are labeled with a cocktail of mouse monoclonal antibodies against known markers for mature human blood cells, such as CD2, CD3, CD14, CD16, CD19, CD24, CD56, CD66b, and glycophorin A. Magnetic iron particles are then attached to the antibodies and lineage-committed cells are depleted in a magnetic field using LD MACS columns. LD columns are recommended for depletion of unwanted cells and good depletion efficiency can be obtained even if the magnetic labeling of the cells is weak. MS and LS columns, designed for positive selection of cells, can also be used for depletion if the magnetic labeling of cells is strong. A flow chart of the protocol steps is presented in Figure 2A.2.2.

BASIC PROTOCOL 2

Materials Mononuclear cells from human cord blood (UNIT 2A.1) Labeling buffer (see recipe), degassed StemSep Human Progenitor Enrichment Kit (StemCell Technologies) containing:

Figure 2A.2.2

Flow chart of the protocol steps to isolate Lin− cells (Basic Protocol 2). Hematopoietic Stem Cells

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Supplement 1

Progenitor Enrichment Cocktail with monoclonal antibodies against CD2, CD3, CD14, CD16, CD19, CD24, CD56, CD66b, and glycophorin A StemSep magnetic colloid MACS LD column (no. 130-042-901, Miltenyi Biotech) Magnetic cell separator MidiMACS (no. 130-042-302, Miltenyi Biotec) 10 ml centrifuge tubes Additional reagents and equipment for preparing mononuclear cell suspension (UNIT 1E.2) Label cells 1. Prepare mononuclear cell suspension (UNIT 2A.1). 2. Add 1 ml of labeling buffer per 8 × 107 mononuclear cells (for fewer cells use 1 ml of labeling buffer). Use degassed buffer only, as the gas bubbles may lead to clogging of the column and decrease the efficacy of immunomagnetic separation. EDTA in the labeling buffer may be replaced by other anticoagulants, such as 0.6% citrate dextrose or citrate phosphate dextrose.

3. Add 100 µl of Progenitor Enrichment Cocktail. 4. Mix well and incubate at room temperature for 15 min. 5. Add 60 µl of StemSep magnetic colloid. 6. Mix well and incubate at room temperature for 15 min.

Perform magnetic separation 7. Place LD column in the magnetic field of the MACS separator. 8. Rinse the column with 2 ml of labeling buffer. 9. Apply labeled cell suspension to the column. If cell aggregates are formed, the cell suspension may be filtered using 30-µm nylon mesh or preseparation filters (no. 130-041-407, Miltenyi Biotech) before applying the cell suspension to the column.

10. Allow unlabeled cells to pass through the column and collect the flowthrough as the Lin− cell fraction. 11. Wash the column two times with 1 ml of labeling buffer (collect in the same tube with the Lin− cells from step 10). 12. Remove the column from the magnetic field and place on a new centrifuge tube. 13. Elute and collect the Lin+ cells two times with 1 ml of labeling buffer. Use the plunger supplied with the column on the second elution. The cells are ready to be used in assays or experiments. The Lin+ cells are often collected for control purposes and used as the positive counterparts for Lin− cells.

REAGENTS AND SOLUTIONS Isolation of Hematopoietic Stem Cells from Human Cord Blood

For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

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Labeling buffer Phosphate-buffered saline (PBS; see recipe) 0.5% (w/v) bovine serum albumin 2 mM disodium EDTA Degas by vacuum Store up to 2 months at 4◦ C Phosphate-buffered saline (PBS) 0.23 g NaH2 PO4 (1.9 mM) 1.15 g Na2 HPO4 (8.1 mM) 9.00 g NaCl (154 mM) Add H2 O to 900 ml Adjust to desired pH (usually 7.2 to 7.4) with 1 M NaOH or 1 M HCl Add H2 O to 1 liter Sterilize by filtering through a 0.22-µm filter or by autoclaving. Store indefinitely at 4◦ C Without adjustment, pH is normally ∼7.3.

COMMENTARY Background Information MACS system was originally developed at the Institute of Genetics, University of Cologne in 1988 to pre-enrich cells for further sorting with flow cytometry. MACS cell separation is based on immunomagnetic selection of labeled cells. In positive selection, mononuclear cells are labeled with paramagnetic microbeads conjugated to specific monoclonal antibodies. Magnetically labeled cells are separated over a column placed in a magnetic field. The labeled cells are retained in the column, while unlabeled cells pass through and can be collected as the unlabeled fraction. The retained labeled cells are eluted from the column after removal from the magnet. Negative selection is a reversed technology, where the unwanted cells are labeled and retained in the column. Unlabeled cells pass through the column and are collected for subsequent applications. The MACS column matrix provides a strong magnetic field to retain cells labeled with minimal amounts of magnetic material. The size of paramagnetic microbeads is small and only a few antigens attached to antibody conjugated microbeads are needed to separate a cell. Therefore, the method is well suited to isolate rare stem cells. Immunomagnetic cell sorting enables fast and gentle separation of viable and highly pure hematopoietic stem cells. The immunomagnetically separated cells can be applied to various downstream applications, such as flow

cytometry, cell culture, colony forming unit assay as well as microscopic, genetic, or molecular analysis.

Critical Parameters and Troubleshooting In order to perform successful immunomagnetic separation for hematopoietic stem cells, one needs to make sure that the mononuclear cell suspension contains no aggregates that can clog the column. Aggregation can be especially problematic when working with cryopreserved cord blood. The mononuclear cells can be resuspended in 200 µl of 1 mg/ml DNaseI before continuing with the labeling. Also, a nylon mesh or commercial preseparation filters may be used to remove cell clumps. Excess thrombocytes in the mononuclear cell population may cause clumping of cells, clog the column, and lower the purity of selected cell populations. The number of thrombocytes may be reduced by additional washing of the mononuclear cell fraction before continuing with the labeling. Use PBS supplemented with 2 mM EDTA or 0.6% citrate for washing and centrifuge for 10 min at 200 × g, 18◦ to 20◦ C. If the antibody bound to the cell is crosslinking, it can form patches, i.e., precipitates of antigen-antibody complexes, resulting in uneven distribution of the antibody on the cell surface. This capping of antibodies can be avoided by using cold solutions and working quickly. However, working on ice is not recommended as it increases the incubation

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Supplement 1

Figure 2A.2.3 Purity of CD34+/− and CD133+/− cell fractions (Basic Protocol 1) as determined by flow cytometry. (A) Representative sample of CD34+ cells shows purity of 97%, and its negative counterpart (CD34− cells) is 99% pure. (B) Representative samples of CD133+ and CD133− cells show purities of 94% and 99%, respectively. SSC, side scatter.

Isolation of Hematopoietic Stem Cells from Human Cord Blood

times. Increased temperature or unnecessary prolonged incubation time may lead to unspecific labeling. Degassing the labeling buffer by vacuum is highly recommended, as the gas can form bubbles in the column matrix and lead to clogging of the column. This lowers the quality of the immunomagnetic separation and may even prevent the elution of the desired cell population. EDTA in the labeling buffer may be replaced by other anticoagulants (for example 0.6% citrate dextrose or citrate phosphate dextrose) if EDTA is expected to hamper subsequent applications such as cell culture. Single column separation of CD34+ or CD133+ cells may result in low purity (typically <50%), yet a fairly large number of cells can be obtained. Two successive column

separations with the additional labeling step (Basic Protocol 1) increase the purity to >90%, but lowers the yield. Highly pure cell fractions are often needed for subsequent applications, so pooling of samples may need to be considered to obtain sufficient cell numbers. Hence, enrichment of hematopoietic stem cells requires balancing between yield and purity, especially when working with a limited source of sample, such as cord blood. The purity of CD34+ , CD133+ , or Lin− cell fractions may be determined by flow cytometry using anti-human CD34, anti-human CD133, or anti-mouse immunoglobulin specific antibodies (as the Lin− cells are depleted using mouse monoclonal antigens), respectively. Typical purities for selected cell fractions are presented in Figures 2A.2.3 and 2A.2.4.

2A.2.8 Supplement 1

Current Protocols in Stem Cell Biology

Figure 2A.2.4 Purity of Lin−/+ cell fractions (Basic Protocol 2) as determined by flow cytometry. Representative sample of Lin− cell fraction is 97% pure. The Lin+ cell fraction is 99% pure. SSC, side scatter.

Anticipated Results

Generally, the recovery of CD34+ , CD133+ , and Lin− cells from cord blood mononuclear cells is 0.86%, 0.21%, and 0.29%, respectively (Kekarainen et al., 2006). The recovery is slightly higher from fresh cord blood when compared to cryopreserved cord blood. The purity of immunomagnetically separated stem cell fractions (CD34+ , CD133+ , and Lin− ) is over 90%, and their control cell populations (CD34− , CD133− and Lin+ ) are nearly 100% pure (Figs. 2A.2.3 and 2A.2.4). The CD34+ and CD133+ cells have similar colony forming potential (84.5 and 80.0 total cfus per 1000 cells, respectively), whereas Lin− cells possess lower capacity to form colonies in CFU assay (57.3 total cfus per 1000 cells).

Time Considerations

Isolation of CD34+ or CD133+ cells (Basic Protocol 1) is performed in ∼1.5 hr and the operation time for isolation of Lin− cells (Basic Protocol 2) is typically 45 min.

Literature Cited Aroviita, P., Teramo, K., Hiilesmaa, V., and Kekomaki, R. 2005. Cord blood hematopoietic progenitor cell concentration and infant sex. Transfusion 45:613-621.

Broxmeyer, H.E., Douglas, G.W., Hangoc, G., Cooper, S., Bard, J., English, D., Arny, M., Thomas, L., and Boyse, E.A. 1989. Human umbilical cord blood as a potential source of transplantable hematopoietic stem/progenitor cells. Proc. Natl. Acad. Sci. U.S.A. 86:3828-3832. Grewal, S.S., Barker, J.N., Davies, S.M., and Wagner, J.E. 2003. Unrelated donor hematopoietic cell transplantation: Marrow or umbilical cord blood? Blood 101:4233-4244. Hao, Q.L., Shah, A.J., Thiemann, F.T., Smogorzewska, E.M., and Crooks, G.M. 1995. A functional comparison of CD34+ CD38− cells in cord blood and bone marrow. Blood 86:3745-3753. Hogan, C.J., Shpall, E.J., McNulty, O., McNiece, I., Dick, J.E., Shultz, L.D., and Keller, G. 1997. Engraftment and development of human CD34(+)enriched cells from umbilical cord blood in NOD/LtSz-scid/scid mice. Blood 90:85-96. Kekarainen, T., Mannelin, S., Laine, J., and Jaatinen, T. 2006. Optimization of immunomagnetic separation for cord blood-derived hematopoietic stem cells. B.M.C. Cell. Biol. 7:30. Vormoor, J., Lapidot, T., Pflumio, F., Risdon, G., Patterson, B., Broxmeyer, H.E., and Dick, J.E. 1994. Immature human cord blood progenitors engraft and proliferate to high levels in severe combined immunodeficient mice. Blood 83:2489-2497.

Hematopoietic Stem Cells

2A.2.9 Current Protocols in Stem Cell Biology

Supplement 1

Isolation of Mesenchymal Stem Cells from Human Cord Blood

UNIT 2A.3

Anita Laitinen1 and Jarmo Laine1 1

Finnish Red Cross Blood Service, Helsinki, Finland

ABSTRACT Cord blood is a rich source of stem cells especially for hematopoietic stem cells. Recently, mesenchymal stem cells (MSCs) have also been shown to exist in cord blood. However, these fibroblast-like multipotent progenitor cells are rather rare in cord blood. Many different methods have been used for their culture. This unit describes one method to obtain MSCs from cord blood and another method to differentiate these cells into osteoblasts, which is one of the lineages that mesenchymal stem cells are capable of differentiating into. The starting material for the protocol is cord blood–derived mononuclear cells. As cord blood contains a great number of erythroid precursors, the glycophorin A–positive cells are depleted using magnetic cell separation to reduce their presence in MSC culture. Osteoblast differentiation and a method to demonstrate the result of the differentiation C 2007 by are also described in this unit. Curr. Protoc. Stem Cell Biol. 1:2A.3.1-2A.3.7.
John Wiley & Sons, Inc. Keywords: cord blood r mesenchymal stem cells r glycophorin A r osteoblasts r von Kossa staining

INTRODUCTION Cord blood is one of the latest sources for mesenchymal stem cells (MSCs). Because MSC populations lack specific cell surface markers many isolation protocols are based on negative selection or rely on culturing of the cells as primary cultures without any other selection methods. MSC isolation is traditionally based on the ability of the cells to adhere to plastic surfaces. Bone marrow (BM) has been the prime source of MSCs. Cord blood differs from BM as the starting material and extra steps are helpful before plating the cells. As cord blood contains a large number of erythroid progenitors that do not necessarily sediment to the bottom layer in Ficoll-Paque density gradient centrifugation, the mononuclear cell fraction can be purified from these contaminating cells by magnetic cell separation (also see UNIT 2A.2). The erythroid progenitor cells express a single-pass transmembrane glycoprotein glycophorin A (GlyA) on their plasma membrane that can be used in cell selection. The GlyA negative (GlyA− ) cells can then be cultured without the confounding presence of erythroid progenitors. This unit presents a protocol for MSC isolation (see Basic Protocol 1) and a protocol for MSC differentiation into osteoblasts (see Basic Protocol 2). The first protocol describes a method for growing MSCs from cord blood mononuclear cells isolated using magnetic cell separation (also see UNIT 2A.2). The second protocol gives a method for differentiating the MSCs into osteoblasts, which is one of the lineages that the MSCs can be differentiated to. The Support Protocol describes a staining method to demonstrate the differentiation of MSCs to osteoblasts. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. Hematopoietic Stem Cells Current Protocols in Stem Cell Biology 2A.3.1-2A.3.7 Published online June 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a03s1 C 2007 John Wiley & Sons, Inc. Copyright


2A.3.1 Supplement 1

NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

ISOLATION OF MESENCHYMAL STEM CELLS FROM HUMAN CORD BLOOD MONONUCLEAR CELLS This protocol describes one method for obtaining MSCs from fresh human cord blood. Mononuclear cells from fresh cord blood isolated using Ficoll-Paque density gradient centrifugation are used as the starting material (UNIT 2A.1). The GlyA− cell selection and the culture of the cells to obtain for MSCs is described in this protocol (also see UNIT 2A.2). The GlyA− cell selection is performed using immunomagnetic selection. The mononuclear cells are labeled with magnetic microbeads attached to an antibody specific to GlyA. The magnetically labeled cells are then depleted in a magnetic field using LD MACS columns. These GlyA− cells are then cultured in conditions that support mesenchymal stem cell growth.

Materials Mononuclear cells from fresh human cord blood (UNIT 2A.1) Labeling buffer (see recipe), degassed Glycophorin A MicroBeads (Miltenyi Biotec) MSC culture medium (see recipe) Phosphate-buffered saline (PBS; see recipe) 0.25% (w/v) trypsin/1 mM EDTA solution Freezing medium: 50% MSC culture medium/40% (v/v) fetal bovine serum/10% (v/v) dimethyl sulfoxide (DMSO), chilled 10-ml and 50-ml centrifuge tubes Centrifuge (preferably swinging-bucket rotor) MACS LD columns (Miltenyi Biotech) Magnetic cell separator (MidiMACS, Miltenyi Biotec) Fibronectin-coated 6-well plates (see recipe) Cryovials Cell-freezing container Liquid nitrogen Additional reagents and equipment for preparing mononuclear cell suspension (UNIT 2A.1) and cell counting (Phelan, 2006) Separate GlyA− cells 1. Prepare mononuclear cell suspension (UNIT 2A.1) in a 50-ml centrifuge tube. Add 80 µl labeling buffer per 107 mononuclear cells (for fewer cells use 80 µl of labeling buffer). Use degassed labeling buffer only, as the gas bubbles may lead to clogging of the column and hamper the immunomagnetic separation. When working with higher cell numbers, scale up the buffer and reagent volumes (e.g., 80 µl of labeling buffer per 107 mononuclear cells or 1600 µl of labeling buffer per 2 × 108 mononuclear cells).

2. Add 20 µl of Glycophorin A MicroBeads per 107 mononuclear cells. Use cold reagents and solutions to avoid capping of antibodies on cell surface.

3. Mix well and incubate at 4◦ to 8◦ C for 15 to 30 min. Isolation of Mesenchymal Stem Cells from Human Cord Blood

4. Add 10 vol of labeling buffer to wash cells/beads. 5. Centrifuge for 10 min at 300 × g, 20◦ C. Discard the supernatant from the wash. 6. Resuspend the cells in 500 µl of labeling buffer per 108 mononuclear cells.

2A.3.2 Supplement 1

Current Protocols in Stem Cell Biology

7. Place the LD column in the magnetic field of the MACS separator. 8. Rinse the column with 2 ml labeling buffer.

Collect GlyA− cells 9. Place a new 10-ml tube under the column and apply the labeled cell suspension to the column. 10. Allow unlabeled cells to pass through the column. The effluent of the unlabeled cell suspension will be clear. The GlyA+ cells remaining in the column gives the red color of the mononuclear cell suspension described in previous steps.

11. Wash the column twice with 1 ml labeling buffer and collect in the same tube with the GlyA− cells from step 10. The total effluent contains the GlyA− cell fraction. If the GlyA+ cell fraction is to be collected remove the column from the magnetic field and place it on a new centrifuge tube. Apply 3 ml of labeling buffer to the column. Use the plunger supplied with the column to flush out the magnetically labeled cell fraction. The color of the eluted cell suspension will be red.

Establish MSC cultures 12. Centrifuge the eluted GlyA− cell fraction for 5 min at 300 × g, room temperature. 13. Resuspend the cells in 5 ml of MSC culture medium and count the cells (Phelan, 2006). 14. Adjust the cells to a concentration of 3.2 × 106 /ml of MSC culture medium. 15. Plate the cells on fibronectin-coated 6-well plates, 3 ml of cell suspension per well (cell density 106 /cm2 ). 16. Incubate the cells overnight. 17. The next day, remove nonadherent cells by changing the medium. The nonadherent cells that are not removed at the first medium change will be removed in subsequent medium changes. Do not wash the plates at this point because the desired cells might not have adhered properly yet.

Maintain MSC cultures 18. Incubate the cells changing the medium twice a week. Monitor the wells for the cell proliferation. The colonies of fibroblast-like cells should be seen within 3 weeks (Fig. 2A.3.1A). If no colonies have appeared within 3 weeks the plates can be discarded.

19. When cell colonies are visible, let the cells grow to 50% to 80% confluency.

Passage cells 20. Wash the wells with PBS. Add 150 µl of trypsin/EDTA solution and incubate at room temperature for 5 min. 21. Neutralize the trypsin with 1 ml of MSC culture medium containing serum and collect the cells into a 50-ml centrifuge tube. Centrifuge for 5 min at 300 × g, room temperature. The cells should loosen easily from the plastic. The cells that are not detached might be unwanted monocyte lineage cells.

22. Replate the cells on 6-well plates at density of 2000 to 3500/cm2 using 3 ml of MSC medium per well.

Hematopoietic Stem Cells

2A.3.3 Current Protocols in Stem Cell Biology

Supplement 1

Figure 2A.3.1 Morphology of the mesenchymal stem cells and von Kossa staining of the osteoblast differentiated cells. (A) Proliferating mesenchymal stem cells from cord blood (100× magnification; Basic Protocol 1). (B) Mesenchymal stem cells differentiated 3 weeks to osteoblasts (100× magnification; Basic Protocol 2). (C) von Kossa stained cells differentiated 3 weeks to osteoblasts (40× magnification; Support Protocol).

At the first passage, if there are only few cells, the cells might be replated to lower density without counting the cells. Just trypsinize the cell colony and plate the cells on a new 6-well plate well or wells.

23. Continue to culture the cells until a sufficient number have been obtained for all necessary experiments or for cryopreservation. 24. Passage the cells when they are ∼80% confluent and if needed change the medium so that the cells will have fresh medium at least twice a week.

Freeze aliquots of MSC 25. To prepare frozen stocks, remove the cells from the plate, centrifuge, and resuspend the cells in chilled freezing medium at 106 cells/ml. 26. Pipet the cell suspension into a cryovial. Place the cryovials into a cell freezing container and place the container in −70◦ C freezer and then later into liquid nitrogen. BASIC PROTOCOL 2

DIFFERENTIATION OF MESENCHYMAL STEM CELLS TO OSTEOBLASTS Osteoblast differentiation of MSCs is used to test their differentiation potential. Osteoblast differentiation is usually verified using von Kossa staining method (see Support Protocol). This protocol describes a basic method to differentiate MSCs to osteoblasts using differentiation medium containing dexamethasone, β-glycerophosphate, and ascorbic acid-2-phosphate, which induce the MSCs differentiation into osteoblasts.

Materials Mesenchymal stem cells (MSCs; Basic Protocol 1) MSC culture medium (see recipe) Differentiation medium (see recipe) 24- or 6- well tissue culture plates Additional reagents and equipment for preparation of mesenchymal stem cell suspension (Basic Protocol 1) and von Kossa staining (Support Protocol) 1. Prepare MSC suspension in MSC culture medium containing 6–10 × 103 cells/ml for 24-well plates or 9.6–16 × 103 cells/ml for 6-well plates. Isolation of Mesenchymal Stem Cells from Human Cord Blood

2. Pipet 1 ml of cell suspension per well for 24-well plate or 3 ml per well for 6-well plate to give a final cell density 3–5 × 103 per cm2 . 3. Let the cells adhere well overnight in tissue culture incubator.

2A.3.4 Supplement 1

Current Protocols in Stem Cell Biology

4. Next day, aspirate the culture medium and add the same volume of differentiation medium to the cells. 5. Change the medium twice a week. 6. Monitor the cell differentiation. The cells will start to form mineralized matrix that is seen as deposit on the cells (Fig. 2A.3.1B).

7. Let the cells differentiate 2 to 5 weeks. 8. Check the differentiation at time points of interest (e.g., 2, 3, 4, and 5 weeks) by von Kossa staining (Support Protocol).

VON KOSSA STAINING FOR OSTEOBLASTS Von Kossa staining is a basic staining method used to demonstrate mineralization by staining the calcium phosphate matrix formed by osteoblasts.

SUPPORT PROTOCOL

Materials Differentiated cells (Basic Protocol 2) Phosphate-buffered saline (PBS; see recipe) 4% (w/v) paraformaldehyde (pH 7.2; see recipe) Deionized water 1% (w/v) silver nitrate 2.5% (w/v) sodium thiosulfate Mirror or aluminum foil UV light source or 60-W lamp 1. Wash the differentiated cells (Basic Protocol 2) with PBS (use 1 ml/well for the 24-well plate and 3 ml/well for the 6-well plate). 2. Fix the cells with 4% paraformaldehyde for 10 min, room temperature. Use 500 µl/well for the 24-well plate and 1 ml/well for the 6-well plate. 3. Aspirate the paraformaldehyde and wash the wells once with PBS and twice with deionized water. 4. Apply 1% silver nitrate solution to the wells (use 500 µl/well for the 24-well plate and 1 ml/well for the 6-well plate). 5. Place the plates on a mirror or aluminum foil and illuminate them with UV light (or use 60-W lamp) for 30 min. The calcium-containing matrix is stained black (Fig. 2A.3.1C).

6. Wash the wells three times for 5 min with deionized water each wash. 7. Apply sodium thiosulfate to the wells (use 500 µl/well for the 24-well plate and 1 ml/well for the 6-well plate) and incubate for 5 min to stop the reaction. 8. Rinse the wells with deionized water. 9. Air dry the cells. 10. Photograph the plates. Semiquantitatively score the staining based on von Kossa staining intensity.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. Current Protocols in Stem Cell Biology

Hematopoietic Stem Cells

2A.3.5 Supplement 1

Differentiation medium α-MEM (Invitrogen) 15% (v/v) fetal bovine serum (Invitrogen) 20 mM HEPES 1× L-glutamine (Invitrogen) 1× penicillin-streptomycin (Invitrogen) 0.1 µM dexamethasone 10 mM β-glycerophosphate 0.05 mM L-ascorbic acid-2-phosphate Prepare fresh medium weekly Store the medium up to 1 week at 4◦ C Fibronectin coating of culture surfaces Prepare 5 ng/ml fibronectin in PBS (see recipe). Pipet 1 ml/well of solution into 6-well plates. Incubate plates for ≥2 hr at 37◦ C. Aspirate the solution before use. The plates can be stored packaged in an air tight container up to 2 weeks at 4◦ C.

Labeling buffer Phosphate-buffered saline (PBS; see recipe) containing: 0.5% (w/v) bovine serum albumin 2 mM disodium EDTA Degas by vacuum Store up to 1 month at 4◦ C MSC culture medium 41% (v/v) DMEM, low glucose (Invitrogen) 40% (v/v) MCDB 201 (Sigma-Aldrich) 15% (v/v) fetal bovine serum (Invitrogen) 1× penicillin-streptomycin (Invitrogen) 1× ITS (insulin-transferrin-selenium) liquid supplement (Sigma-Aldrich) 1× linoleic acid–BSA (Sigma-Aldrich) 5 × 10−8 M dexamethasone (Sigma-Aldrich) 0.1 mM L-ascorbic acid-2-phosphate (Sigma-Aldrich) Store up to 1 week at 4◦ C Paraformaldehyde, 4% (w/v), pH 7.2 Dissolve 20 g paraformaldehyde in ∼450 ml of PBS (see recipe). Heat the mixture to ∼70◦ C while stirring. Add 1 M NaOH until the solution clarifies. Cool and adjust the pH to 7.2 using 1 M HCl. Adjust the volume to 500 ml with phosphate-buffered saline (PBS, see recipe). Divide into 10-ml aliquots and store up to 3 months at −20◦ C, thaw just before use.

Phosphate-buffered saline (PBS)

Isolation of Mesenchymal Stem Cells from Human Cord Blood

0.23 g NaH2 PO4 (1.9 mM) 1.15 g Na2 HPO4 (8.1 mM) 9.00 g NaCl (154 mM) Add H2 O to 900 ml Adjust to desired pH (usually 7.2 to 7.4) with 1 M NaOH or 1 M HCl, if necessary Add H2 O to 1 liter Sterilize by filtering through 0.22-µm filter or autoclaving Store indefinitely at 4◦ C Store up to 1 month at 4◦ C

2A.3.6 Supplement 1

Current Protocols in Stem Cell Biology

COMMENTARY Background Information Isolation of mesenchymal stem cells (MSCs) was initially described from mouse bone marrow (Friedenstein et al., 1976). MSCs have been isolated from different tissues including adipose tissue, peripheral blood, and cord blood. MSCs can be isolated from cord blood using many different protocols. The MSCs from cord blood are typically isolated without any cell selection method using mononuclear cells as starting material. However, as cord blood contains a large number of erythroid progenitors, their potential confounding effect on MSC isolation can be avoided by magnetic cell separation of GlyA+ cells. The method described here is based on the protocol to isolate multipotent adult progenitor cells (MAPCs) from bone marrow as described by Reyes et al. (2001). The protocol to isolate MAPCs from cord blood was tested, but instead of MAPCs it gave rise to MSCs. The protocol was further developed to give a better yield of MSC. Serum concentration was 15%, which is much higher than has been used to isolate MAPCs from bone marrow. In lower serum concentration the efficiency rate for growing MSCs from cord blood proved to be only 11%. MSCs are capable of differentiating into mesenchymal cell types like osteoblasts, chondroblasts, and adipocytes (Pittenger et al., 1999). The osteoblast differentiation occurs under appropriate tissue culture conditions demonstrated by Jaiswal et al. (1997). The reagents supporting osteoblast differentiation in vitro are dexamethasone, β-glycerophosphate, and ascorbic acid-2phosphate. The osteoblast differentiation of the cells has traditionally been demonstrated by so-called von Kossa staining, in which the mineralized calcium matrix of the osteoblasts is stained with silver nitrate.

Critical Parameters and Troubleshooting As the MSCs are very rare in cord blood and they tend to adhere to plastic, it is important to work fast or use siliconized glassware whenever possible. The time between the collection

of cord blood and starting the culture procedure should be kept as short as possible. To get better yield of MSCs from cord blood, the cord blood units should be processed within 15 hr (Bieback et al., 2004). All work must be carried out aseptically.

Anticipated Results

The number of GlyA− cells obtained from one cord blood unit varies between 1 × 108 and 3 × 108 . The efficiency to grow mesenchymal stem cells from fresh cord blood using the described protocol is ∼40% to 50%. In practice, a single cord blood unit may give rise to 1 to 2 MSC colonies that can be expanded and cultured for 5 to 20 passages.

Time Considerations

The isolation of GlyA− cells is performed in ∼1.5 hr. Setting up the culture of GlyA− cells takes ∼45 min including preparation of medium. Obtaining MSC colonies is a very slow process that usually takes between 8 and 21 days.

Literature Cited Bieback, K., Kern, S., Kluter, H., and Eichler, H. 2004. Critical parameters for the isolation of mesenchymal stem cells from umbilical cord blood. Stem Cells 22:625-634. Friedenstein, A.J., Gorskaja, J.F., and Kulagina, N.N. 1976. Fibroblast precursors in normal and irradiated mouse hematopoietic organs. Exp. Hematol. 4:267-274. Jaiswal, N., Haynesworth, S.E., Caplan, A.I., and Bruder, S.P. 1997. Osteogenic differentiation of purified, culture-expanded human mesenchymal stem cells in vitro. J. Cell Biochem. 64:295-312. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A3F.1-A.3F.18. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., and Marshak, D.R. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284:143147. Reyes, M., Lund, T., Lenvik, T., Aguiar, D., Koodie, L., and Verfaillie, C.M. 2001. Purification and ex vivo expansion of postnatal human marrow mesodermal progenitor cells. Blood 98:26152625.

Hematopoietic Stem Cells

2A.3.7 Current Protocols in Stem Cell Biology

Supplement 1

Isolation and Assessment of Long-Term Reconstituting Hematopoietic Stem Cells from Adult Mouse Bone Marrow

UNIT 2A.4

David Kent,1 Brad Dykstra,1 and Connie Eaves1 1

University of British Columbia, Vancouver, Canada

ABSTRACT Suspensions of multipotent hematopoietic stem cells with long-term repopulating activity can now be routinely isolated from adult mouse bone marrow at purities of 30%. A robust method for obtaining these cells in a single step using multiparameter cell sorting to isolate the CD45mid lin− Rho− SP subset is described here, together with a detailed protocol for assessing their regenerative activity in mice transplanted with single cells. These procedures provide unprecedented power and precision for characterizing the molecular and biological properties of cells with hematopoietic stem cell activity at the C 2007 by John Wiley single cell level. Curr. Protoc. Stem Cell Biol. 3:2A.4.1-2A.4.23.
& Sons, Inc. Keywords: stem cell isolation r flow cytometry r hematopoiesis r single cell transplants r multi-lineage reconstitution

INTRODUCTION The hematopoietic system is responsible for producing all of the various types of specialized blood cells in numbers required throughout life. These specialized cells comprise those of the lymphoid lineages, which include B cells, T cells, and natural killer cells, as well as a diversity of myeloid cell types, including erythrocytes and monocytes, neutrophilic, eosinophilic and basophilic granulocytes, and megakaryocytes, which generate platelets. The mature forms of most of these cells cannot proliferate and also have a short lifespan (a few days or weeks). Thus, in order for the required numbers of blood cells to be maintained, new blood cells must be continuously generated. Much evidence indicates that this is achieved throughout adult life by a multistep differentiation process—one that originates in a rare population of cells individually capable of generating all of the differentiated blood cell types. At each division, these very primitive cells must also be able to produce at least one daughter cell that is similarly competent but remains undifferentiated (i.e., to execute a self-renewal division; reviewed in Bryder et al., 2006). These properties of multipotency and self-renewal ability are the basis for designating such cells as hematopoietic stem cells (HSCs). Identifying the molecular elements that determine the properties of the progeny of HSCs, as well as the elements that may alter these outcomes over successive divisions, are longterm goals of the field. They are also essential to the development of better therapies for many diseases where normal blood cell production is limiting or defective. Investigating the mechanisms that regulate HSC functions requires methods for their specific detection, quantification, and purification. This has historically posed a major challenge because of the extremely low frequency of HSCs in hematopoietic tissue (less than 10–4 ) and the inability of HSCs to be distinguished morphologically from other types of primitive cells. Furthermore, no unique molecular signature of HSCs has been identified; indeed, it is not yet clear that all HSCs can even be captured by a common set of molecular features. Somatic Stem Cells Current Protocols in Stem Cell Biology 2A.4.1-2A.4.23 Published online November 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a04s3 C 2007 John Wiley & Sons, Inc. Copyright


2A.4.1 Supplement 3

Therefore, at present, the most specific and reliable methods for identifying HSCs make use of retrospective functional assays that detect the ability of HSCs to produce mature blood cells for prolonged periods of time in vivo (4 to 6 months in mice, Fig. 2A.4.1). This involves injecting the test cells into recipient mice whose own ability to produce new blood cells has been severely compromised, but whose survival has been independently assured either by coinjection of radioprotective cells or through the use of sublethally irradiated W-deficient recipients (Szilvassy et al., 1989, 1990; Jones et al., 1990). At a time point 4 to 6 months later, the circulating blood cells in these recipient mice are analyzed to determine whether they include myeloid and lymphoid cells generated from the originally injected test cells. Over the years, a variety of strategies have been utilized to distinguish donor-derived cells in the recipient mice. These include DNA-based detection of sex-mismatched transplants by Southern blot (Szilvassy et al., 1990), PCR or FISH analysis (Ramshaw et al., 1995), electrophoretic detection of different glucose isomerase isoforms (Trevisan and Iscove, 1995) or hemoglobin allotypes (Harrison, 1980), and, more recently, flow cytometric detection of a cell surface alloantigen (CD45; Spangrude et al., 1988) or the product of a reporter transgene like green fluorescent protein (GFP; Wagers et al., 2002). A major advantage of the latter strategies is the ease with which they can be combined with flow cytometric identification of different mature blood cell types, thus allowing the differentiation potential of the injected cells to be tracked with considerable precision and sensitivity.

Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

Figure 2A.4.1 In vivo transplantation assay to detect HSCs. HSCs can be retrospectively identified by their ability to produce multiple WBC types for an extended period of time after being transplanted into irradiated recipients. This provides a method for functionally assessing the hallmark properties of HSCs (i.e., self-renewal and multipotentiality). One or more test cells that are congenic for a single cell surface marker (CD45) are injected into irradiated recipients. After a minimum of 16 weeks, a blood sample is collected and then evaluated by flow cytometry for the presence of donor-derived WBCs. If donor-derived WBCs of both myeloid and lymphoid lineages are detected, the original test cells are inferred to have contained a HSC.

2A.4.2 Supplement 3

Current Protocols in Stem Cell Biology

If a test cell population is transplanted together with competitor cells with predefined hematopoietic activity, the relative competitive reconstituting ability of the test cells can be measured based on the proportion of donor-derived cells subsequently detected in the blood or other hematopoietic organs. This qualitative measure of HSC activity in the test cells can then be used to estimate the number of HSCs in the test cell population. However, such calculations are limited by the requisite assumption (unless validated) of constancy in the average mature cell output per injected HSC (Harrison, 1980; Harrison et al., 1993; Audet et al., 2001). Injection of the test cells at doses that become limiting for long-term reconstitution allow HSC frequencies to be calculated in the absence of any assumption about their mature blood cell output, except for the requirement of a certain predetermined threshold level and composition (Szilvassy et al., 1990). Table 2A.4.1 summarizes the criteria that have been used for defining HSC activity in various studies. It also illustrates how this definition has evolved over time. During the last decade, various methods have been developed for isolating populations of cells from suspensions of adult mouse bone marrow that represent >20% pure HSCs (Osawa et al., 1996; Adolfsson et al., 2001; Christensen and Weissman, 2001; Chen et al., 2002; Benveniste et al., 2003; Uchida et al., 2003; Matsuzaki et al., 2004; Kiel et al., 2005; Balazs et al., 2006; Wagers and Weissman, 2006). However, it has now been established that many of these methods rely on the use of markers that change according to the activation/cycling status of HSCs (Rebel et al., 1996; Sato et al., 1999; Tajima et al., 2001; Uchida et al., 2004; Zhang and Lodish, 2005). Therefore, most purification strategies are applicable only for steady-state bone marrow in which most adult HSCs are normally quiescent (Harrison and Lerner, 1991; Bradford et al., 1997; Cheshier et al., 1999). Consequently, these methods are often not useful for the direct enumeration of HSCs that have been biologically or genetically manipulated, and functional endpoints of HSC activity remain the gold standard for their detection. Interestingly, it has recently been shown that the CD150+ CD48− phenotype of HSCs found in adult bone marrow (Kiel et al., 2005) extends to cytokine-mobilized and regenerating HSCs as well as HSCs in the fetal liver (Yilmaz et al., 2006). Although the specificity of this phenotype for HSCs manipulated in vitro remains to be examined, these studies suggest that more direct readouts for these cells may be possible. In this unit, we describe a robust method for the isolation of HSCs from the bone marrow of normal adult C57Bl/6 mice which consistently yields a suspension that is ∼30% pure based on the frequency of cells that will continue to reconstitute irradiated recipient mice with both lymphoid and myeloid progeny for at least 4 months (see Basic Protocol 1; Uchida et al., 2003; Dykstra et al., 2006). This method of HSC purification involves isolating a very rare subset of all bone marrow cells (0.004%) that are distinguished by their expression of intermediate levels of CD45 and undetectable levels of various “lineage” (lin) markers (Spangrude et al., 1988) in combination with an ability to efflux both rhodamine 123 (Rho; Bertoncello et al., 1991) and Hoechst 33342 (Hst; Goodell et al., 1996). These cells are thus referred to as CD45mid lin− Rho− SP (side population) cells. Such levels of purity allow transplants of single cells to replace limiting dilution methods for enumerating HSC directly from assessments of positively repopulated recipients. Importantly, however, we have found that the methods used to detect the in vivo progeny of the injected cells and the criteria used to define a positively repopulated mouse are key to determining the types of input cells that are identified. Therefore, in addition to outlining a protocol for performing single-cell transplants, we have included a description of a procedure for analyzing the donor-derived white blood cells (WBCs) that are subsequently generated, as well as the criteria that we have found useful for discriminating HSC-repopulated mice (Dykstra et al., 2007).

Somatic Stem Cells

2A.4.3 Current Protocols in Stem Cell Biology

Supplement 3

Table 2A.4.1 Definitions of Long-Term Multi-Lineage Repopulation by Donor-Derived Cells

Publication

Hosta

Competitor cells

Min. % repopulation

Lineages

Weeks posttransplantation

Szilvassy et al. (1990)

8.5 Gy female B6/C3

1-2 × 105 compromised

Detectable by Southern, ∼5%

Male DNA in BM and thymus

5 to 10

Morrison and Weissman (1994)

Split 9-9.2 Gy B6 2 × 105

0.3% of each lineage

Ly5.2, B220+Mac1, CD3+Gr1

16+

Rebel et al. (1994)

9-9.5 Gy B6

2 × 105 compromised or 4 × 105 Sca-

20%

None required

11 to 15

Trevisan et al. (1996)

3 Gy W41 /W41

None

“Detectable”, ∼0.1-1%

Erythroid (via GPI-shift assay)

32 to 52

Osawa et al. (1996) 9.5 Gy B6

500 CD34lowKSL “Detectable”

Mac1/Gr1, Thy1/B220 (not required)

12 +survival

Miller and Eaves (1997)

9 Gy B6 or 4 Gy W41 /W41

105 or nothing

1%

Gr-1+, SSClow

16+

Sudo et al. (2000)

9.5 Gy B6

2 × 105

1%

Gr1/Mac1, CD4/CD8, B220

12

Ema et al. (2000)

9.5 Gy B6

2 × 105

1%

Gr1/Mac1, CD4/CD8, B220

12

Adolfsson et al. (2001)

9.5 Gy B6

1.5-2 × 105

0.5% of total, 0.1% myeloid

Gr1/Mac1

16

Szilvassy et al. (2003)

Split 9 Gy B6

2 × 105

5%

Gr1/Mac1, Thy1, B220

5,10,17,26

Uchida et al. (2003) 4 Gy W41 /W41

None

0.05% of each lineage

B220, CD5, Gr1/Mac1

16+

de Haan et al. (2003)

10 Gy B6

2 × 105

2% of each lineage

Gr1/Mac1, Thy1, B220

8

Matsuzaki et al. (2004)

10.5 Gy B6

2 × 105

1%

Gr1/Mac1, B220/CD3

12

Kiel et al. (2005)

Split 11 Gy B6

2 × 105

Above background (0.1%-0.3%)

Gr1/Mac1, B220, 16 CD3

Ema et al. (2005)

9.5 Gy B6

2 × 105

1% of each lineage

Gr1/Mac1, CD4/CD8, B220

12 to 16

Zhang and Lodish (2005)

Split 10 Gy B6

1-2 × 105

1%

Thy1, B220, Mac1, Gr1, Ter119

16

Rossi et al. (2005)

Split 9.2 Gy B6

3 × 105

“Detectable”

Mac1, B220, TCRβ+

28

None

1% of total at 16 wk, 1% of each lineage at any time

Gr1/Mac1, B220/CD5

4,8,12,16

Dykstra et al. (2006) 4 Gy W41 /W41

a B6 = C57B1/6J mice; Gy = total irradiation in units of Gray. Sources and types of irradiation vary. The qualifier “split” is used for groups that split the irradiation dose into two different blocks of time in order to allow recovery of the hematopoietic system between doses.

2A.4.4 Supplement 3

Current Protocols in Stem Cell Biology

ISOLATION OF CD45mid lin− Rho− SP CELLS FROM ADULT MOUSE BONE MARROW

BASIC PROTOCOL 1

Since the advent of high-speed, multicolor flow cytometry, many groups interested in the biology of HSCs have exploited this technology to isolate these rare cells as relatively pure populations. The technique described here is attractive because of its simplicity and ability to generate cells that show minimal to no antibody labeling, which can reduce HSC engraftment efficiency (Szilvassy et al., 1989; Gilner et al., 2007). Rather, it combines the removal of cells labeled with fluorescent antibodies against non-HSCs with the positive selection of cells that can exclude the retention of two fluorescent dyes (Hst and Rho).

Materials Mice (strain C57B1/6J, 8 to 12 weeks old) HBSS/2% FBS: Hanks’ balanced salt solution (HBSS; StemCell Technologies) containing 2% (w/v) fetal bovine serum (FBS; StemCell Technologies) 0.8% (w/v) ammonium chloride (NH4 Cl) solution (StemCell Technologies), ice cold 0.1 mg/ml DNase I (Sigma) Serum-free medium (SFM; see recipe, or purchase StemSpan serum-free expansion medium from StemCell Technologies, cat. no. 09600) 20 µg/ml rhodamine 123 (Rho; Molecular Probes) in sterile PBS 500 µg/ml Hoechst 33342 (Hst; Sigma) in sterile PBS 1 µg/ml reserpine (Sigma) in DMSO HBSS/2% FBS plus 1.2 µl mouse FcR blocking antibody (2.4G2, hybridoma available from ATCC; Ab available commercially from StemCell Technologies, cat. no. 01504) Lin antibody cocktail, biotinylated (StemSep Hematopoietic Progenitor Enrichment Cocktail, StemCell Technologies) CD45-allophycocyanin (CD45-APC, Becton Dickinson) Streptavidin-phycoerythrin (streptavidin-PE, Becton Dickinson) HBSS/2% FBS (both reagents available from StemCell Technologies) plus 1 µg/ml propidium iodide (PI; Sigma) Dissecting instruments including scissors and forceps, sterile 21-G, 1-in. general-use sterile hypodermic needles (Becton Dickinson, cat. no. 305165) Tabletop centrifuge Flow cytometer sample tubes (e.g., Becton Dickinson, cat. no. 352058 or 352063) 37◦ C water bath with cover, of sufficient depth to ensure that the water is above the level of liquid in the tubes incubated 50-ml conical tubes (e.g.: Becton Dickinson, cat. no. 352070) 5-ml filter-capped tubes (Becton Dickinson, cat. no. 358235) Flow cytometer with sorting capability (e.g., FACSCalibur; Becton Dickinson) 96-well U-bottom plates (e.g., Nunc, cat. no. 163320; optional, necessary if performing single-cell isolation as in Basic Protocol 2) Additional reagents and equipment for flow cytometry (Robinson et al., 2007) Prepare the cells 1. Harvest the bone marrow cells from femurs and tibia of C57B1/6J mice by flushing the interior of the bones with a 21-G, 1-in. needle using 3 ml of HBSS/2% FBS. 2. Lyse the red blood cells (RBCs) by adding ∼10 ml of ice cold 0.8% NH4 Cl solution, then incubating the nucleated cells on ice for exactly 10 min, vortexing lightly after the first 5 min. Somatic Stem Cells

2A.4.5 Current Protocols in Stem Cell Biology

Supplement 3

3. Centrifuge the cell suspension 5 min at 300 × g, 4◦ C. At this time a cell count may be performed using an ∼1:10 dilution, which should show a recovery of between 5 and 6 × 107 cells from both tibias and both femurs of each mouse. If a preliminary enrichment step is desired (e.g., to reduce reagent costs and sorting time when isolating HSCs from bone marrow harvests from multiple mice), an immunomagnetic depletion step can be used (e.g., using the biotinylated StemSep Hematopoietic Progenitor Enrichment Cocktail and EasySep technology available from StemCell Technologies: see http://www.stemcell.com/technical/19756-PIS.pdf).

4. Remove the supernatant, add 50 µl DNase I (to minimize cell clumping), flick the pellet to disperse the cells, and resuspend the cells in serum-free medium (SFM) to a concentration of 107 per ml. 5. Dispense 100-µl aliquots of this suspension into each of three control flow cytometer sample tubes (PI-only, APC-only, PE-only) on ice. Cells for the Rho, Hst, or reserpine controls DO NOT need to be removed until later (steps 11 and 12). It is recommended to have a PI-only control and a reserpine control to define the SP gate, as well as single-stain controls, with PI, for APC, PE, and Rho.

Prepare cells for Rho and Hst incubations 6. Dilute the remaining cells in SFM to 2 × 106 per ml in a 50-ml conical tube. If the required volume is greater than 50 ml, all the cells should be stained with Rho in one 50-ml tube, then diluted to 2 × 106 per ml in additional 50-ml tubes. For example, 2 × 108 cells would need to be diluted in 100 ml of medium. Thus, in step 7, 50 ml of cells in medium and 50 ml of medium alone should be warmed. In step 8, 500 µl of 20 µg/ml Rho would be added to the tube containing the cells. 12.5 ml of this suspension would then be distributed into three new 50-ml tubes (leaving 12.5 in the original tube). Next, 12.5 ml of warm medium would be added to each of the four tubes.

7. Warm the cells at 37◦ C for 15 min in a water bath. 8. Add 50 µl of 20 µg/ml Rho per 10 ml of cells (a 1:200 dilution) to the 50-ml conical tube with the cells. Mix well. For incubation and washing, the maximum recommended volume is 25 ml per tube. Thus, if the starting population contains more than 5 × 107 cells, first divide the cells evenly among multiple 50-ml conical tubes (see annotation to step 6 above).

9. Incubate the cells for exactly 30 min in the dark in the 37◦ C water bath, ensuring that the water level is above the level of the liquid in the tube(s) containing the cells. IMPORTANT NOTE: Longer or shorter incubations will decrease the yield and/or purity of HSCs in the final cell suspension to be sorted. Also, it is important to keep the temperature at a constant 37◦ C (i.e., do not put other items into the water bath during the incubation).

10. Fill each tube with HBSS/2% FBS. Centrifuge 5 min at 300 × g, 4◦ C, and decant the supernatant. Add 100 µl of 0.1 mg/ml DNase I and resuspend the cells. 11. Add warm (37◦ C) SFM to the cells to give a concentration of 2 × 106 cells per ml. If cells are in more than one tube, combine all cells into a single 50-ml conical tube. This will ensure that all cells are equally exposed to the Hst dye. Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

12. Remove 0.5 ml for the Rho-only control to be incubated alongside the sample in step 16. 13. Add 100 µl of 500 µg/ml Hst per 10 ml of cells (a 1:100 dilution) to the 50-ml tube containing the remaining cells.

2A.4.6 Supplement 3

Current Protocols in Stem Cell Biology

14. Set up the reserpine control by removing 100 µl from the 50-ml tube and adding 1 µl of 1 µg/ml reserpine (to this 100-µl aliquot only) for incubation in parallel with the other samples. Mix well. Reserpine is toxic to the cells and will result in the detection of a higher percentage of dead (PI+ ) cells when these are later analyzed by flow cytometry.

15. Distribute the Rho- and Hst-treated cells from the 50-ml tube among multiple 50-ml conical tubes with a maximum of 25 ml per tube, as described in step 8. 16. Incubate the following tubes for 90 min in the dark in a 37◦ C water bath, as in step 9: a. x number of tubes (25 ml maximum each, from step 15) with the main sample evenly distributed. b. The Rho-only control (from step 12). c. The reserpine-treated control (from step 14). At the end of the incubation, place the Rho-only control and the reserpine-treated controls on ice and in the dark, along with the controls set aside in step 5. 17. At the end of the incubation period, fill the 50-ml tubes containing Rho/Hst-treated cells with ice-cold HBSS/2% FBS and place them on ice for 5 min. IMPORTANT NOTE: From this point onwards, everything must be on ice and in the dark; precautions include turning off the lights while the cells are being manipulated.

18. Centrifuge the cells 5 min at 300 × g, 4◦ C, remove the supernatant, add 100 µl of 0.1 mg/ml DNase I to the cell pellet, and flick to resuspend the cells.

Stain with antibody 19. Dilute the cells to 107 cells per ml in ice-cold HBSS/2% FBS plus 1.2 µl of 2.4G2 (blocking reagent) per ml. 20. Combine the cells into a single 50-ml conical tube and incubate them on ice and in the dark for another 10 min. 21. Add the following antibodies to the following tubes: a. To the control tube “PE-only” (see step 5) add 0.5 µl lin biotinylated antibody cocktail to prepare the “lin-PE” control tube. b. To the control tube “APC-only” (see step 5) add 0.5 µl CD45-APC. c. To the main (sample) tube (see step 20; this can still be in the original 50-ml conical tube) add 5 µl/ml of the lin biotinylated cocktail and 5 µl/ml CD45-APC. 22. Mix the cells in each tube and incubate the tubes on ice in the dark for ∼30 min. 23. Transfer the cells from the main (sample) tube (step 21, substep c) to a smaller (5-ml) tube. Wash the cells in this tube as well as those in the tubes from step 21, substeps a and b, by filling each tube to 5 ml with ice-cold HBSS/2% FBS, centrifuging 5 min at 300 × g, 4◦ C, and removing the supernatants. For the control tubes (step 21, substeps a and b), resuspend the pellets in 100 µl HBSS/2% FBS. For the main (sample) tube (step 21, substep c), add 50 µl of 0.1 mg/ml DNase I to the cell pellet, and flick to resuspend the cells. The clumping that the DNase I acts to prevent, results from lysed cells produced by the centrifugation steps, hence the repeated DNase treatments.

24. Add sufficient HBSS/2% FBS to the cells in the main (sample) tube for a final concentration of 107 cells/ml.

Somatic Stem Cells

2A.4.7 Current Protocols in Stem Cell Biology

Supplement 3

25. Add 0.1 µl streptavidin-PE to the lin-PE control tube (after resuspending the cells to 100 µl). 26. Add 1 µl/ml streptavidin-PE to the main (sample) tube. 27. Mix the cells in each tube and incubate the tubes on ice in the dark for ∼15 min. 28. Wash all tubes by adding 3 ml ice-cold HBSS/2% FBS, centrifuging 5 min at 300 × g, 4◦ C, and removing the supernatants. 29. To each control tube, add 200 µl of 1 µg/ml PI/HBSS/2% FBS. 30. To the sample tube, add 50 µl DNase I, then add 200 µl of 1 µg/ml PI and transfer to a 5-ml filter-capped tube. Add 200 µl of the 1 µg/ml PI solution to the original tube to rinse out all of the remaining cells and transfer to the filter-capped tube. Repeat this washing of the original tube until the total volume in the filter-capped tube is ∼1.2 to 1.5 ml.

Sort cells 31. Use a PI-only control to set the single WBC and viable gates (Fig. 2A.4.2A,B). Robinson et al. (2007) contains detailed protocols in flow cytometry.

32. Use the single marker-stained controls (Rho, PE, APC) to set up the compensation for each detection channel to be used during the sort and apply these settings to the sample tube.

Figure 2A.4.2 Flow cytometric profiles for isolating CD45mid lin− Rho− SP cells. Stained bone marrow cells were gated first using FSC/SSC (A) and PI (B) to exclude debris, erythrocytes, dead cells, and cell clumps. Gates were then set around the CD45mid (C), SP (D), and lin− Rho− (E) populations. (F) In combination, these five gates select for ∼0.004% of the original bone marrow cells.

2A.4.8 Supplement 3

Current Protocols in Stem Cell Biology

33. Establish the SP gate by using a reserpine (or verapamil) control to highlight the area where the SP cells in the untreated sample appear. The voltages of the UV-red and UV-blue channel should be set such that the main population (upper right, Figure 2A.4.2, panel D) is centered at approximately two-thirds of the x and y axes to ensure good resolution of the SP tip.

34. Set gates, as shown in Figure 2A.4.2, and sort the cells into a desired volume of medium (e.g., one cell in 100 to 200 µl medium per well of a 96-well U-bottom, for Basic Protocol 2).

TRANSPLANTATION OF SINGLE CELLS AND TRACKING OF THEIR REGENERATED WBC CELL PROGENY

BASIC PROTOCOL 2

The most direct method of assessing the in vivo regenerative activity of individual HSCs is to transplant them as single cells into irradiated but radioprotected recipients and then follow over time the numbers and types of progeny WBCs produced. The use of sublethally irradiated C57BL6-W41 /W41 (W41 /W41 ) mice as recipients allows this type of experiment to be performed in a simple fashion with the same specificity and sensitivity as is achieved using lethally irradiated wild-type recipients given a minimally competing transplant of long-term repopulating cells also containing sufficient short-term repopulating cells to ensure the survival of the host (Miller et al., 1996; Audet et al., 2001; Uchida et al., 2003). The W41 /W41 mouse has a c-kit tyrosine kinase receptor with reduced signaling activity, and this results in the generation of a 17-fold smaller pool of HSCs in the marrow of adult mice of this genotype (Miller et al., 1996). Consequently, a sublethal dose of radiation is sufficient to create a permissive environment for normal HSCs that are transplanted into such recipients, and extensive donor chimerism can be achieved from transplants of single cells without the need to coinject any other cells. Here, we describe a method for the initial visualization and injection of single cells into sublethally irradiated W41 /W41 recipients, as well as a method for analyzing their mature progeny subsequently detectable in the circulation. This protocol is described for an experiment in which a single cell has been sorted into 100 to 200 µl of medium in each well of a 96-well plate. Each single cell is then visually confirmed, harvested into a syringe, and transplanted into a mouse. The method can also be scaled for larger cell numbers or modified for other types of manipulations.

Materials Single HSCs sorted into 100 to 200 µl medium in a 96-well plate (Basic Protocol 1) Irradiated recipient mice (strain W41/W41 or C57B1/6J, >5 weeks old; see annotation to step 6 of this protocol regarding irradiation), exposed for 2 to 3 min to a heat lamp 0.8% (w/v) ammonium chloride (NH4 Cl) solution (StemCell Technologies), ice cold HBSS/2% FBS: Hanks’ balanced salt solution (HBSS; StemCell Technologies) containing 2% (w/v)fetal bovine serum (FBS, StemCell Technologies) 2× blocking reagent (see recipe) Antibody cocktails for peripheral blood analysis (see recipe) HBSS/2% FBS plus 1 µg/ml propidium iodide (PI; Sigma) Tabletop centrifuge with microtiter plate carrier Inverted microscope (preferably with movable stage) Insulin syringes with 28-G, 0.5-in needles (Becton Dickinson) Heparinized capillary tubes (e.g., Fisher) 12 × 75–cm tubes with caps (Becton Dickinson cat. no. 352057) 96-well U-bottom microtiter plates (e.g., Nunc, cat. no. 163320)

Somatic Stem Cells

2A.4.9 Current Protocols in Stem Cell Biology

Supplement 3

ScreenMates 1.4-ml round-bottom storage tubes in snap rack (Thermo Scientific, cat. no. 4246) Flow cytometer (equipped with a HeNe and argon laser, e.g.: Becton Dickinson FACSCalibur) Additional reagents and equipment for flow cytometry (Robinson et al., 2007) Inject single cells 1. Centrifuge the 96-well plate containing the HSCs 5 min at ∼180 × g, 4◦ C, to bring the cells to the bottom of each well without damaging them. 2. Visualize each cell using a standard inverted microscope. Cells from the CD45mid lin− Rho− SP subset appear as small round cells with a crisp border when the focus is slightly altered.

3. Once the well is confirmed to have one and only one cell, mark it and proceed to the next well. When the desired number of single cells have been identified (this should not take more than 30 min), place the entire plate on ice. 4. For each well, fill a single-use insulin syringe (with 28-G, 0.5-in. needle) with ∼300 µl PBS and remove all air bubbles. IMPORTANT NOTE: It is critical to remove all air bubbles for the following operations.

5. Using the syringe, gently push ∼50 µl of the 300 µl into the well to dislodge the cell from the bottom of the well (this must be done with care in order to avoid causing any liquid to overflow the well). Next, use the syringe to remove almost all of the liquid from the well and then gently dispense it back into the well. Finally, aspirate all of the liquid from the well into the syringe, being very careful not to create any air bubbles. 6. Immediately inject the entire volume into the tail vein of an irradiated mouse that has just been exposed to an infra-red heat lamp for ∼2 to 3 min. Alternatively, the filled syringes may be placed in a beaker inside a container of ice until the injections are completed. Depending on the available strains of mice, a lethal or sublethal dose of irradiation should be used. C57BL/6J mice, for example, require a lethal dose (900 cGy) and should receive additional (but genetically distinct) cells to ensure their radioprotection (e.g., of the same genotype as the host animal); mice homozygous for the W41 allele can be irradiated with a sublethal dose (400 cGy) and then do not require cotransplantation of additional cells for their survival.

Analyze peripheral blood Ensure that additional blood samples are taken for the appropriate positive and negative controls for the Ly5.1 and Ly5.2 antibodies. If the experiment utilizes C57BL/6J (Ly5.2) donors and W41 /W41 Ly5.1 recipients, then use a peripheral blood sample from a C57BL/6J mouse as a positive control for the donor cells and a peripheral blood sample from a noninjected W41 /W41 mouse as a negative control (i.e., no donor cells; see steps 13 and 23 for special instructions on how to use these samples). Use the remaining cells from the positive and negative controls for the single-stained and PI-only controls, as appropriate (see Table 2A.4.2). Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

7. Collect 50 to 75 µl of blood from each mouse (using, e.g., tail-vein, retro-orbital sinus, saphenous vein, or cheek-pouch bleed) into heparinized capillary tubes. 8. Flush the blood sample into a 12 × 75–cm tube. Keep all samples on ice following collection.

2A.4.10 Supplement 3

Current Protocols in Stem Cell Biology

Table 2A.4.2 Set up of 96-Well Plates for Staining

Well

Cells

Additions

A1

+ or – control cells

PIa

A2

+ control cells (Ly5.2)

PI + Ly5.2 FITC single-stain control

A3

– control cells (Ly5.1)

PI + Ly5.1 APC single-stain control

A4

+ or – control cells

PI + Ly6g/Mac1 PE single-stain control

A5

+ or – control cells

PI + B220 PE single-stain control

A6

+ or – control cells

PI + Ly1 PE single-stain control

B1

– control cells (Ly5.1)

PI + GM cocktailb

B2

– control cells (Ly5.1)

PI + B cell cocktailb

B3

– control cells (Ly5.1)

PI + T cell cocktailb

B4

+ control cells (Ly5.2)

PI + GM cocktailb

B5

+ control cells (Ly5.2)

PI + B cell cocktailb

B6

+ control cells (Ly5.2)

PI + T cell cocktailb

C1

Mouse 1

PI + GM cocktailb

C2

Mouse 1

PI + B cell cocktailb

C3

Mouse 1

PI + T cell cocktailb

C4

Mouse 2

PI + GM cocktailb

C5

Mouse 2

PI + B cell cocktailb

C6c

Mouse 2

PI + T cell cocktailb

a PI = 1 µg/ml propidium iodide in HBSS/2% FBS. b See recipe for antibody cocktails for peripheral blood analysis. c After well C6, the remainder of the plate can be filled with additional samples from new mice, with three wells needed to analyze each mouse.

9. To lyse the RBCs, add 2 ml of ice-cold 0.8% NH4 Cl solution and vortex the suspension lightly. 10. Incubate for exactly 10 min on ice, vortexing lightly at the 5-min mark. It has been our experience that a longer lysis step leads to significant loss of granulocytes. In order to minimize this loss, it is important to perform the lysis step on ice and only for 10 min in total.

11. Add 5 ml of HBSS/2% FBS and centrifuge the cells 5 min at 300 × g, 4◦ C. 12. Remove the supernatant, leaving no more than 50 µl. 13. Add 150 µl of 2× blocking reagent to each sample tube and 300 µl of 2× blocking reagent to the positive and negative control tubes. The extra amount in each control will be necessary to stain the positive and the negative controls with each of the antibodies to be used.

14. Incubate ∼10 min at room temperature or 20 min on ice. To save time, it is useful to set up the plate for staining during this incubation period.

15. Using a multichannel pipettor, aliquot 3 µl of each antibody cocktail into the appropriate wells of a 96-well plate and then add 50 µl of cells into each well (be sure to add antibody before cells). An efficient way to set up a 96-well plate is shown in Table 2A.4.2.

Somatic Stem Cells

2A.4.11 Current Protocols in Stem Cell Biology

Supplement 3

16. Incubate the cells for ≥30 min on ice. It is critical that the cells be incubated at least 30 min on ice.

17. Add 150 µl of 0.8% NH4 Cl to each well in order to perform an additional RBC lysis. Better RBC lysis results in more efficient flow cytometric acquisition and cleaner profiles. Use of a plastic reagent trough and a multichannel pipettor for this step will save time.

18. Centrifuge the plate(s) 5 min at 300 × g, 4◦ C. 19. Remove the supernatant from each well with a Pasteur pipet attached to a vacuum source, or quickly flick the entire plate over the sink to remove the supernatant. Flicking the plate is faster, but cell recoveries will be reduced and some cell pellets may be completely lost if the plate is flicked too violently.

20. Place the 1.4-ml round-bottom tubes in the specially designed 96-slot snap rack. If there is an automated plate reader for the flow cytometer (e.g., a High-Throughput System, HTS, from Becton Dickinson), omit step 20 and proceed directly to step 21 without transferring the cells.

Figure 2A.4.3 Flow cytometric profiles of WBCs from a mouse that is highly reconstituted with transplanted cells. Viable (A) WBCs (B) are shown in panel (C) with the donor antigen (Ly5.2) on the x axis and the recipient antigen (Ly5.1) on the y axis. Panels (D-L) include cells only within singly stained Ly5.1 or Ly5.2 gates (to exclude any cell doublets). Myeloid cells are Ly6g/Mac1+ (D), granulocytes are Ly6g+ /SSChigh (E, F), B cells are B220+ (G), and T cells are Ly1+ (H). Panels (I-L) show recipient versus donor contributions to the myeloid (I), granulocyte (J), B cell (K) and T cell (L) compartments. Panel (F) is a replicate plot of panel (E), but shown as a contour plot with an easily defined boundary to allow the high SSC gate to be drawn. Note the dominance of donor-derived cells in all three lineages, particularly the myeloid lineage.

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Current Protocols in Stem Cell Biology

Figure 2A.4.4 Flow cytometric profiles of WBCs from a mouse showing a weak and lineage-restricted pattern of transplant-derived reconstitution. Viable (A) WBCs (B) are shown in panel (C) with the donor antigen (Ly5.2) on the x axis and the recipient antigen (Ly5.1) on the y axis. Panels (D-L) include cells only within singly stained Ly5.1 or Ly5.2 gates (to exclude any cell doublets). Myeloid cells are Ly6g/Mac1+ (D), granulocytes are Ly6g+ /SSChigh (E, F), B cells are B220+ (G) and T cells are Ly1+ (H). Panels (I-L) show recipient versus donor contributions to the myeloid (I), granulocyte (J), B cell (K) and T cell (L) compartments. Panel (F) is a replicate plot of panel (E), but shown as a contour plot with an easily defined boundary to allow the high SSC gate to be drawn. Note that most of the donor-derived cells are T cells.

21. Add 100 µl of 1 µg/ml PI in HBSS/2% FBS into each well and then transfer all of the contents directly into the corresponding 1.4-ml plastic tubes using a multichannel pipettor. If using the HTS system, resuspend the cells in 100 µl of 1 µg/ml PI in HBSS/2% FBS and leave them in the plate.

Acquire cells on flow cytometer 22. Use the PI-only control to set the viable and single-marker-stained WBC gates (Fig. 2A.4.2A,B). 23. Use the single-marker-stained controls (FITC, PE, APC) to set up the compensation required for each channel to be used and apply these settings to the sample tube(s). 24. Run the positive and negative control samples to verify that the settings are correct, and to assist with subsequent analysis of the remaining samples. Figures 2A.4.3 to 2A.4.5 depict representative plots for mice that show strong repopulation (Fig. 2A.4.3), lineage-restricted repopulation (Fig. 2A.4.4), and undetectable levels of repopulation (Fig. 2A.4.5).

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Figure 2A.4.5 Flow cytometric profiles of WBCs from a mouse showing no transplant-derived reconstitution. Viable (A) WBCs (B) are shown in panel (C) with the donor antigen (Ly5.2) on the x axis and the recipient antigen (Ly5.1) on the y axis. Panels (D-L) include cells only within the singly stained Ly5.1 or Ly5.2 gates (to exclude any doublets). Myeloid cells are Ly6g/Mac1+ (D), granulocytes are Ly6g+ /SSChigh (E, F), B cells are B220+ (G) and T cells are Ly1+ (H). Panels (I-L) show recipient versus donor contributions to the myeloid (I), granulocyte (J), B cell (K) and T cell (L) compartments. Panel (F) is a replicate plot of panel (E), but shown as a contour plot with an easily defined boundary to allow the high SSC gate to be drawn. Note the dominance of host-derived cells in all three lineages.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Antibody cocktails for peripheral blood analysis B cell peripheral blood analysis cocktail: Titrate and combine the following antibodies, diluted for the addition of 3 µl of the antibody cocktail to 50 µl of cells: Ly5.2-FITC (eBiosciences) Ly5.1-APC (eBiosciences) B220-PE (Becton Dickinson)

Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

Granulocyte/monocyte (GM) peripheral blood analysis cocktail: Titrate and combine the following antibodies, diluted for the addition of 3 µl of the antibody cocktail to 50 µl of cells: Ly5.2-FITC (eBiosciences) Ly5.1-APC (eBiosciences) Mac-1-PE (Becton Dickinson) Ly6g-PE (Becton Dickinson) continued

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T cell peripheral blood analysis cocktail: Titrate and combine the following antibodies, diluted for the addition of 3 µl of the antibody cocktail to 50 µl of cells: Ly5.2-FITC (eBiosciences) Ly5.1-APC (eBiosciences) Ly1-PE (Becton Dickinson) Store cocktails at 4◦ C, observing manufacturer’s expiration dates. Recipes for antibody cocktails should be individually calculated based on titrations of the antibody components. Adjust amounts of antibody as necessary when new antibodies are titrated and use sterile PBS to make up the remainder of the volume in each case.

Blocking reagent, 2x 1 ml rat serum (Sigma) 50 µl mouse FcR blocking antibody (2.4G2 hybridoma available from ATCC; Ab available commercially from StemCell Technologies, cat. no. 01504) 9.495 ml Hanks’ balanced salt solution (HBSS; StemCell Technologies) Store up to 8 weeks at 4◦ C Serum-free medium (SFM) To prepare 100 ml: 77 ml Iscove’s Modified Dulbecco’s Medium (IMDM, StemCell Technologies) 20 ml BIT serum substitute (mixture of bovine serum albumin, insulin, and transferrin; StemCell Technologies) 1 ml 10–2 M 2-mercaptoethanol in H2 O 1 ml 2 mM L-glutamine in IMDM 1 ml penicillin/streptomycin solution (StemCell Technologies; contains 100 U/ml penicillin and 100 µg/ml streptomycin) Prepare fresh and keep cold Alternatively, StemSpan serum-free expansion medium may be purchased from StemCell Technologies.

COMMENTARY Background Information In the early 1950s, cellular extracts prepared from the bone marrow or spleen of mice were found to be protective against lethal doses of radiation in mice (Lorenz et al., 1951). For the next few years, it was hotly debated as to whether this protective effect was mediated by a humoral factor or by transplantable cells with regenerative activity. By the mid-1950s, the use of transplants of cytogenetically marked donor cells resolved this issue by demonstrating the ability of protective transplants to take over the new blood supply of the host (Ford et al., 1956). The spleen colony assay, introduced by Till and McCulloch (1961), was the first method for quantifying cells with multilineage reconstituting activity, and the cells identified were called colony-forming units– spleen (CFU-S). Use of the CFU-S assay to characterize the properties of the cells thus identified and their regulation allowed this group and others to formulate many of the ba-

sic concepts of HSC biology, including those covered by the terms of self-renewal, multipotentiality, lineage restriction, and differentiation. The demonstration of heterogeneity among CFU-S was also documented, and the concept of a pre-CFU-S cell was suggested (Hodgson and Bradley, 1979; Schofield and Dexter, 1985). However, many years elapsed before convincing experimental evidence of a distinct population of this latter type was obtained (Ploemacher and Brons, 1989; Jones et al., 1990). This came from the use of Rho staining and counterflow centrifugal elutriation to separate CFU-S from cells with more durable repopulating activity. Almost simultaneously, retroviral marking experiments provided definitive evidence of the presence in normal adult bone marrow of multipotent self-renewing hematopoietic cells with lifelong reconstituting ability (LTRCs; Dick et al., 1985; Keller et al., 1985; Lemischka et al., 1986).

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Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

Advances in multiparameter flow cytometry led to the development of more sophisticated methods for isolating rare subsets of cells including primitive hematopoietic cells. A strategy for removing the more prevalent maturing populations using a panel of cell surface markers expressed as part of their terminal differentiation program (the so-called lin markers) was introduced together with the positive selection of cells expressing Sca-1 (Spangrude et al., 1988). This methodology was subsequently refined by the addition of antibodies to c-kit as part of the positive selection strategy (Okada et al., 1991). Nevertheless, both the lin− Sca-1+ and lin− Sca1− c-kit+ (KSL) populations were shown to contain CFU-S as well as LTRCs, suggesting persisting functional heterogeneity within the KSL subset. Later experiments that made use of a variety of markers confirmed this prediction. These experiments included testing antibodies to other cell surface markers such as CD34 (Osawa et al., 1996), CD27 (Wiesmann et al., 2000), flk2/flt3 (Adolfsson et al., 2001; Christensen and Weissman, 2001), endoglin/CD105 (Chen et al., 2002), the signaling lymphocyte activation molecule (SLAM) family receptors CD150 and CD244 (Kiel et al., 2005), endothelial cell protein C receptor (EPCR)/CD201 (Balazs et al., 2006), and α-2 integrin/CD49b (Wagers), and/or examining differences in their staining with Rho (Bertoncello et al., 1991; Wolf et al., 1993; Benveniste et al., 2003) or Hst (Goodell et al., 1996; Majolino et al., 1997; Uchida et al., 2003; Matsuzaki et al., 2004). Cells with an ability to efflux Hst are often visualized using two emission wavelengths to allow the SP phenotype to be identified (Goodell et al., 1996). The sorting strategy described here exploits this latter approach by combining it with lin+ cell removal and coselection of adult mouse bone marrow cells that can also efflux Rho efficiently (Uchida et al., 2003). The measurement of long-term multilineage donor reconstitution ability is central to identifying HSC activity in the in vivo posttransplant setting. However, it is important to note that the definition of exactly what is longterm multi-lineage reconstitution has evolved over the years. With improvements in the antibodies that are now commercially available, as well as in flow cytometry equipment and analysis software, immune-based quantification of the frequency of different types of donor-derived WBCs has become the norm. In addition, the availability of congenic mice expressing immunologically distinguishable

forms of CD45 and monoclonal antibodies raised against these alloantigens (CD45.1 = Ly5.1 and CD45.2 = Ly5.2) has permitted convenient and effective coincident distinction of their donor or host origin. Table 2A.4.1 summarizes the criteria used to define HSC activity in various studies and demonstrates the evolution of the endpoints used to infer the presence of an HSC in the original transplant.

Critical Parameters Careful alignment, calibration, and maintenance of the flow cytometer machine used to isolate rare cells is of utmost importance for the successful and reproducible isolation of HSCs at high purity. This is particularly true for isolating purified HSC populations based on their CD45mid lin− Rho− SP phenotype. In particular, visualization of the SP fraction is very sensitive to how the UV laser is calibrated. On some instruments, it may be necessary to sort the cells at a reduced speed (i.e., <2000 cells per second) to ensure that the phenotype collected is consistent. During laser setup, it is useful to ensure tight signals with calibration beads and to make sure that the positive and negative controls show a good separation of the populations to be distinguished. It is also advisable to keep all sample tubes at 4◦ C if the sort is anticipated to take more than 15 min. Additionally, it is advisable to validate the accuracy and efficiency of any single-cell deposition step to ensure that single cells and only single cells are indeed deposited into each well. Consistent staining and incubation periods, especially with the Rho and Hst dyes, are also critical. Even if the cell sorter is optimally configured, a good sort result is unlikely if the staining has been inconsistent or incomplete. Single-stain, fluorescence minus one (FMO; see Table 2A.4.3), and negative controls are useful indicators of whether one can proceed with confidence. For the analyses of cells in the peripheral blood of transplanted mice, titration of each antibody reagent to select for an appropriate concentration is critical (regardless of the information provided by the supplier). Subsequently, it is advisable to make large batches of each antibody cocktail (e.g., sufficient for analysis of 100 to 200 samples) to decrease interand intra-experimental variability. When analyzing samples, it is also important to use the control samples to establish the correct voltage and compensation settings before running the test samples, and to keep these settings consistent for all of the samples.

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When analyzing multiple lineages of donor-derived cells, several potential pitfalls need to be considered. Figures 2A.4.3 to 2A.4.5 serve to illustrate some of these. Each of these figures shows data from a single mouse that was transplanted 16 weeks before with a single CD45mid lin− Rho− SP cell resulting in high, multi-lineage repopulation (Fig. 2A.4.3), low, lineage-restricted repopulation (Fig. 2A.4.4), and no repopulation (Fig. 2A.4.5), respectively. All gates drawn are the same for each mouse with the viable cells (PI− ) and the WBCs gated in panels A and B. The first thing to note is that in all cases, a population of WBCs that represent ∼0.3% to 0.4% of the total events seen appears to be positive for both Ly5.1 and Ly5.2 (C). These events represent cell doublets and need to be excluded by appropriate gating before any further analyses are performed. To determine the extent to which the donor-derived WBCs include lymphoid and myeloid elements also requires that the lineage-specific gating strategy be sufficiently stringent to ensure that any false positives are completely excluded. This is of particular relevance to establishing whether donor-derived myeloid cells are present which, due to their short lifespan, allow short-term repopulation to be discriminated from longterm reconstitution with the greatest sensitivity. Because a subpopulation of CD3+ T cells can bind low levels of anti-Gr-1 (RB68C5; Bryder et al., 2004), use of this antibody to infer the presence of myeloid cells can be problematic. In the protocol described here, we therefore utilize a different monoclonal antibody (clone 1A8) that is more specific to Ly6G (Fleming et al., 1993). To provide further specificity in the detection of granulocytes, their high light side-scattering characteristics (SSC) can be examined. In Figures 2A.4.3 to 2A.4.5, we have shown the GMhigh SSChigh population in both dot plot format and as a contour plot (F and G). Underneath each of the lineages, we have also shown whether the cells costained with CD45.1 or CD45.2, using the donor and recipient gates that were originally drawn. This serves to illustrate the contribution of each donor-derived clone to each particular WBC lineage evaluated. Finally, Figure 2A.4.6 shows how, even within the gate for (Ly6g/Mac1)+ cells, confirmation of an association with granulocytes using the SSChigh gate can be very useful.

Troubleshooting See Tables 2A.4.3 and 2A.3.4 for guides to possible causes and solutions to problems

encountered in cell isolation and staining experiments.

Anticipated Results Isolation of cells from the mouse bone marrow For normal adult mice, suspensions obtained from the bone marrow should be cloudy and pinkish. Following the RBC lysis step and centrifugation of the nucleated cells, the supernatant should be clear and a strong whitish-red pellet with red edges (and sometimes streaks through the white pellet) should be clearly visible. Resuspension of the cell pellet in a small volume (50 µl) of DNase I is advised for the optimal generation of a single-cell suspension from such initial pellets. Preparation of cells for flow cytometry The final suspension should be contained in a volume of ∼1.5 ml and should be filtered before sorting or analysis using a filter-topped tube to remove any residual clumps of cells. The cell suspension should also be transparent or translucent (dependent on the cell concentration) when resuspended in HBSS/2% FBS. Flow cytometric plots for isolating the CD45mid lin− Rho− SP cells Figure 2A.4.2 shows representative flow cytometric plots for a typical CD45mid lin− Rho− SP cell sorting procedure. First, the viable WBCs are gated based on their FSC/SSC properties and their exclusion of PI (panel B). The next three plots (panels C to E) show the viable cells as they appear in a plot of CD45 staining versus FSC (panel C), the UV blue versus UV red fluorescence (panel D), and lin staining versus Rho fluorescence (panel E). The SP gate (panel D) is drawn based on a reserpine control which inhibits the transporter responsible for Hst dye efflux (Zhou et al., 2001). The plot in panel F shows the UV plot when it is gated for a CD45mid lin− Rho− phenotype. The final population represents ∼0.004% of the bone marrow cells from a normal adult C57Bl/6J mouse. Flow cytometric plots for GM, B cell, and T cell repopulation Figures 2A.4.3 to 2A.4.5 are representative flow cytometric plots for donor-derived clones that are multi-lineage (Fig. 2A.4.3), lineage-restricted (Fig. 2A.4.4), or absent (Fig. 2A.4.5). Careful gating, as described in the Critical Parameters section, should give a robust assessment of whether or not a mouse is repopulated with donor-derived cells in multiple WBC lineages.

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Figure 2A.4.6 Careful gating is required to convincingly demonstrate multi-lineage WBC reconstitution with transplant-derived cells. Viable (A) WBCs (B) are shown with the donor antigen (Ly5.1) on the x axis and the granulocyte-macrophage markers (Ly6g and Mac1) on the y axis. When cells in these two panels are back-gated in the SSC versus FSC plots (C, D, respectively), the high-SSC properties of the highly Ly6g/Mac1+ cells and the low-SSC properties of cells showing intermediate levels of Ly6g/Mac1 positivity can be seen. To focus on the capture of granulocytes, it is helpful to use a Ly6g+ /SSChigh gate, as described in Figures 2A.4.3 to 2A.4.5. In the above case, this would reveal an absence of detectable donor-derived Ly6g+ /SSChigh cells.

Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

Time course analysis of peripheral blood samples Various patterns of reconstitution can be discerned when the number and types of donor-derived WBCs present in the blood of transplanted mice are measured at different times over the initial 24 weeks posttransplantation (Dykstra et al., 2007). For example, the progeny of short-term repopulating cells are typically seen after 4 and 8 weeks, but not thereafter. If more than one cell is transplanted, then the pattern obtained is likely to represent some combination of the patterns that each individual repopulating cell would have contributed.

Time Considerations The time required to harvest the bone marrow cells will vary depending on the number of HSCs desired, but should not take longer than 10 to 15 min per mouse assuming that the marrow from both femurs and both tibias are harvested. The staining procedure takes ∼5 hr, including the three lengthy incubations needed (90, 30, and 30 min). Time spent on the cell sorter will also vary depending on the number of cells desired and whether these need to be collected individually or in bulk. In our experience, ∼200 to 400 CD45mid lin− Rho− SP cells can be routinely isolated on a FACSVantage in ∼2.0 hr. When single-cell transplants are

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Table 2A.4.3 Troubleshooting Guide for the Isolation of CD45mid lin− Rho− SP Cells from Normal Adult Mouse Bone Marrow

Problem

Possible cause

Solution

Insufficient time on the cell sorter to obtain the desired cell number

Not enough input cells; too many contaminating cells.

Harvest more bone marrow and consider a pre-sort immunomagnetic lin+ depletion step (e.g., using EasySep, StemCell Technologies). This will debulk the sample, but the absolute yields will also decrease when adding this step.

All cells are Rho+

The Rho incubation was too long (this is a very sensitive incubation and even 5 min longer will cause a significant difference)

A full time-course analysis may need to be done to determine the proper concentration and incubation length in any given lab. Three populations should be seen: one that is Rho– (∼the lowest 10%), one that is Rhodull (∼the next lowest 40%) and one that is Rhobright (∼the highest 50%). To correctly set the gates, look at the flow cytometry plots in contours to visualize these three populations.

There is no SP in the Hst-stained cells

The incubation was too long.

A full time-course analysis may need to be done to determine the proper concentration and incubation time. If the stain is done in SFM, an SP population that is ∼1% to 2% of the total bone marrow is expected. The reserpine control is necessary to allow the true SP population to be defined.

The inhibitory drug was accidentally added to the “test” cells.

Be very careful with media, pipet tips, and drug addition

The UV laser was poorly aligned

The SP population is not always stable and can appear different on different machines

Drug was not added

Be very careful with media, pipet tips, and drug addition

Cells were all dead

Reserpine is toxic to cells and it may be necessary to lower the concentration. Cell viability in this control should be between 40% and 80%.

Drug no longer has activity

Try a new batch or lot of the drug

Negative population in the sorted sample looks to be more fluorescent than in the unstained control

Sometimes, when using multiple colors, cells can appear autofluorescent. Typically this is caused by the other fluorochromes in the tube.

Use a fluorescence minus one (FMO) control to set the negative gates for each stain. This is prepared by preparing a tube with all of the fluorochromes except the one for which the negative population is being defined (Roederer, 2001).

The cell pellet does not resuspend after staining

Too much volume was added before When resuspending cells, it is most efficient to flick the resuspending pellet in a small volume of medium. This prevents clumps of cells from breaking off and floating in suspension.

All of the cells are PI+

All of the cells are dead

Reserpine control did not work

Do not leave cells overnight unless the medium is supplemented with 10%-50% serum and this is done before the cells are purified. Note that purified HSCs in serum have a propensity to differentiate fairly rapidly. It would also help to check reagents and lab tools for potential hazards to cells (e.g., bleach).

The fluorochrome in PE is very Check the single stained controls to ensure bright and stains the vast majority compensation values are set correctly of cells, and is spilling over into the PI detection channel

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Table 2A.4.4 Troubleshooting Guide for the Detection of Single CD45mid lin− Rho− SP Cells Isolated by Flow Cytometry from Normal Adult Mouse Bone Marrow

Problem

Possible cause

Cannot find the single The cells are not there cells in individual wells The medium is full of debris and cells cannot be distinguished The focal plane is not correct

The bottom of the well is scratched and nothing can be distinguished More than one cell is present

Mice die before the first peripheral blood analysis time point

The granulocyte population of the stained peripheral blood represents less than 5% of the total

A clear separation is not apparent between a specific antibody and the negative control

Irradiation dose was too high

Not enough radioprotective cells were coinjected Excessive lysis of granulocytes occurred during the RBC lysis step

The mouse’s granulocyte compartment is low The antibodies were not titrated properly

Solution Check the settings and calibration on the flow cytometer used. Test beads will often give better results than cells. There are a variety of adjustments that can be made to the sort settings (check with the flow operator or technical representative). Filter the medium before sorting into it with a 0.22-µm filter. This will remove most debris. Adjust the field of view so the entire well is in view, then use the coarse adjustment to focus on the center of the well. This is typically where the cell will be and will establish the approximate viewing plane. It also helps to sort 100-1000 viable cells of any phenotype into the first well to figure out the correct plane. Recommended plates are Nunc 96-well U-bottom plates (see Basic Protocol 1 materials list). Check the settings and calibration on the flow cytometer used. Test beads will often give better results than cells. There are a variety of adjustments that can be made to the sort settings (check with the flow operator or technical rep). Ensure that only 1 cell (and no more than 1.5 droplets) were deposited into each well. Sometimes cells can stick together, though this is rare for the CD45mid lin– Rho− SP phenotype. Finally, this may be attributable to focusing on air bubbles or debris in the medium. Recalibrate the irradiation equipment, perform an LD90 on the strain of mice that is being used, and lower the irradiation dose accordingly. Ensure that the helper cell dose (if used) is accurate and sufficient for the strain of mouse being used. Shorten the RBC lysis period, do not incubate on a shaker or tube rocker, and keep on ice.

Restain the mouse’s peripheral blood without the lysis step to confirm this or do a blood smear to check for granulocytes. These antibodies should all give clear populations of positive cells; retitrate at the same concentration of cells.

The voltage may be too low

The T cell or B cell population looks different between mouse samples

Reanalyze the controls and confirm that the correct compensation values are set. Different amounts of antibody Be very careful when distributing the antibody cocktail into cocktail were added to each individual wells. Ensure that the pipettor accurately dispenses well the desired volume. One of the mice is sick. When mice are afflicted with a disease or infection, the T cell and/or B cell populations may vary from normal.

It is a good idea to re-bleed and reanalyze such mice a week or two later to confirm this if no outward physical signs of sickness are apparent. A blood smear could also be performed to assess cell types present in a given mouse.

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to be performed, the cells should be allowed to settle and be identified prior to starting the injections. Nevertheless, it should be possible to complete these tasks within 1 to 3 hr, again depending on the scale of the experiment. Thus, an entire experiment can be performed within a period of 12 hr. It is not advisable to leave cells for more than 4 to 5 hr before injecting them into irradiated hosts, unless a culture period is part of the experiment. The length of time required to perform the peripheral blood analyses on transplanted mice is more highly dependent on the number of mice to be analyzed. The collection of the blood samples can take between 30 min (∼10 mice) and 3 hr (∼100 mice). The staining steps (using directly labeled antibodies) take between 1 and 2.5 hr, and the flow cytometric analyses take between 30 min and 2.5 hr. Thus overall, analysis of 10 to 100 samples can be readily completed within a single working day.

Literature Cited Adolfsson, J., Borge, O.J., Bryder, D., TheilgaardMonch, K., Astrand-Grundstrom, I., Sitnicka, E., Sasaki, Y., and Jacobsen, S.E.W. 2001. Upregulation of flt3 expression within the bone marrow Lin– Sca1+ c-kit+ stem cell compartment is accompanied by loss of self-renewal capacity. Immunity 15:659-669. Audet, J., Miller, C.L., Rose-John, S., Piret, J., and Eaves, C.J. 2001. Distinct role of gp130 activation in promoting self-renewal divisions by mitogenically stimulated murine hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 98:1757-1762. Balazs, A.B., Fabian, A.J., Esmon, C.T., and Mulligan, R.C. 2006. Endothelial protein C receptor (CD201) explicitly identifies hematopoietic stem cells in murine bone marrow. Blood 107:2317-2321. Benveniste, P., Cantin, C., Hyam, D., and Iscove, N.N. 2003. Hematopoietic stem cells engraft in mice with absolute efficiency. Nat. Immunol. 4:708-713. Bertoncello, I., Bradley, T.R., Hodgson, G.S., and Dunlop, J.M. 1991. The resolution, enrichment and organization of normal bone marrow high proliferative potential colony forming cell subsets on the basis of rhodamine-123 fluorescence. Exp. Hematol. 19:174. Bradford, G.B., Williams, B., Rossi, R., and Bertoncello, I. 1997. Quiescence, cycling, and turnover in the primitive hematopoietic stem cell compartment. Exp. Hematol. 25:445-453. Bryder, D., Sasaki, Y., Borge, O.J., and Jacobsen, S.E. 2004. Deceptive multilineage reconstitution analysis of mice transplanted with hemopoietic stem cells, and implications for assessment of stem cell numbers and lineage potentials. J. Immunol. 172:1548-1552.

Bryder, D., Rossi, D.J., and Weissman, I.L. 2006. Hematopoietic stem cells: The paradigmatic tissue-specific stem cell. Am. J. Pathol. 169:338346. Chen, C.Z., Li, M., de Graaf, D., Monti, S., Gottgens, B., Sanchez, M.J., Lander, E.S., Golub, T.R., Green, A.R., and Lodish, H.F. 2002. Identification of endoglin as a functional marker that defines long-term repopulating hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 99:15468-15473. Cheshier, S.H., Morrison, S.J., Liao, X., and Weissman, I.L. 1999. In vivo proliferation and cell cycle kinetics of long-term self-renewing hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 96:3120-3125. Christensen, J.L. and Weissman, I.L. 2001. Flk-2 is a marker in hematopoietic stem cell differentiation: A simple method to isolate long-term stem cells. Proc. Natl. Acad. Sci. U.S.A. 98:1454114546. de Haan, G., Weersing, E., Dontje, B., van Os, R., Bystrykh, L.V., Vellenga, E., and Miller, G. 2003. In vitro generation of long-term repopulating hematopoietic stem cells by fibroblast growth factor-1. Dev. Cell. 4:241-251. Dick, J.E., Magli, M.C., Huszar, D., Phillips, R.A., and Bernstein, A. 1985. Introduction of a selectable gene into primitive stem cells capable of long-term reconstitution of the hemopoietic system of W/Wv mice. Cell 42:71-79. Dykstra, B., Ramunas, J., Kent, D., McCaffrey, L., Szumsky, E., Kelly, L., Farn, K., Blaylock, A., Eaves, C., and Jervis, E. 2006. High-resolution video monitoring of hematopoietic stem cells cultured in single-cell arrays identifies new features of self-renewal. Proc. Natl. Acad. Sci. U.S.A. 103:8185-8190. Dykstra, B., Kent, D., Bowie, M., McCaffrey, L., Hamilton, M., Lyons, K., Lee, S., Brinkman, R., and Eaves, C. 2007. Long-term propagation of distinct hematopoietic differentiation programs in vivo. Cell Stem Cell 1:218-229. Ema, H., Takano, H., Sudo, K., and Nakauchi, H. 2000. In vitro self-renewal division of hematopoietic stem cells. J. Exp. Med. 192:1281-1288. Ema, H., Sudo, K., Seita, J., Matsubara, A., Morita, Y., Osawa, M., Takatsu, K., Takaki, S., and Nakauchi, H. 2005. Quantification of selfrenewal capacity in single hematopoietic stem cells from normal and Lnk-deficient mice. Dev. Cell. 8:907-914. Fleming, T.J., Fleming, M.L., and Malek, T.R. 1993. Selective expression of Ly-6G on myeloid lineage cells in mouse bone marrow: RB68C5 mAb to granulocyte-differentiation antigen (Gr-1) detects members of the Ly-6 family. J. Immunol. 151:2399-2408. Ford, C.E., Hamerton, J.L., Barnes, D.W.H., and Loutit, J.F. 1956. Cytological identification of radiation chimaeras. Nature 177:452-454. Gilner, J.B., Walton, W.G., Gush, K., and Kirby, S.L. 2007. Antibodies to stem cell marker

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antigens reduce engraftment of hematopoietic stem cells. Stem Cells 25:279-288. Goodell, M.A., Brose, K., Paradis, G., Conner, A.S., and Mulligan, R.C. 1996. Isolation and functional properties of murine hematopoietic stem cells that are replicating in vivo. J. Exp. Med. 183:1797-1806. Harrison, D.E. 1980. Competitive repopulation: A new assay for long-term stem cell functional capacity. Blood 55:77-81. Harrison, D.E. and Lerner, C.P. 1991. Most primitive hematopoietic stem cells are stimulated to cycle rapidly after treatment with 5-fluorouracil. Blood 78:1237-1240. Harrison, D.E., Jordan, C.T., Zhong, R.K., and Astle, C.M. 1993. Primitive hemopoietic stem cells: Direct assay of most productive populations by competitive repopulation with simple binomial, correlation and covariance calculations. Exp. Hematol. 21:206-219. Hodgson, G.S. and Bradley, T.R. 1979. Properties of hematopoietic stem cells surviving 5-fluorouracil treatment: Evidence for a preCFU-S cell? Nature 281:381-382. Jones, R.J., Wagner, J.E., Celano, P., Zicha, M.S., and Sharkis, S.J. 1990. Separation of pluripotent haematopoietic stem cells from spleen colonyforming cells. Nature 347:188-189. Keller, G., Paige, C., Gilboa, E., and Wagner, E.F. 1985. Expression of a foreign gene in myeloid and lymphoid cells derived from multipotent haematopoietic precursors. Nature 318:149154. Kiel, M.J., Yilmaz, O.H., Iwashita, T., Yilmaz, O.H., Terhorst, C., and Morrison, S.J. 2005. SLAM family receptors distinguish hematopoietic stem and progenitor cells and reveal endothelial niches for stem cells. Cell 121:11091121.

Isolation and Assessment of Long-Term Reconstituting Hematopoietic Cells

Miller, C.L., Rebel, V.I., Lemieux, M.E., Helgason, C.D., Lansdorp, P.M., and Eaves, C.J. 1996. Studies of W mutant mice provide evidence for alternate mechanisms capable of activating hematopoietic stem cells. Exp. Hematol. 24:185194. Morrison, S.J. and Weissman, I.L. 1994. The longterm repopulating subset of hematopoietic stem cells is deterministic and isolatable by phenotype. Immunity 1:661-673. Okada, S., Nakauchi, H., Nagayoshi, K., Nishikawa, S., Nishikawa, S.I., Miura, Y., and Suda, T. 1991. Enrichment and characterization of murine hematopoietic stem cells that express c-kit molecule. Blood 78:1706-1712. Osawa, M., Hanada, K.I., Hamada, H., and Nakauchi, H. 1996. Long-term lymphohematopoietic reconstitution by a single CD34low/negative hematopoietic stem cell. Science 273:242-245. Ploemacher, R.E. and Brons, R.H.C. 1989. Separation of CFU-S from primitive cells responsible for reconstitution of the bone marrow hemopoietic stem cell compartment following irradiation: Evidence for a pre-CFU-S cell. Exp. Hematol. 17:263-266. Ramshaw, H.S., Rao, S.S., Crittenden, R.B., Peters, S.O., Weier, H.U., and Quesenberry, P.J. 1995. Engraftment of bone marrow cells into normal unprepared hosts: Effects of 5-fluorouracil and cell cycle status. Blood 86:924-929. Rebel, V.I., Dragowska, W., Eaves, C.J., Humphries, R.K., and Lansdorp, P.M. 1994. Amplification of Sca-1+ Lin- WGA+ cells in serum-free cultures containing steel factor, interleukin-6, and erythropoietin with maintenance of cells with long-term in vivo reconstituting potential. Blood 83:128-136.

Lemischka, I.R., Raulet, D.H., and Mulligan, R.C. 1986. Developmental potential and dynamic behavior of hematopoietic stem cells. Cell 45:917927.

Rebel, V.I., Miller, C.L., Thornbury, G.R., Dragowska, W.H., Eaves, C.J., and Lansdorp, P.M. 1996. A comparison of long-term repopulating hematopoietic stem cells in fetal liver and adult bone marrow from the mouse. Exp. Hematol. 24:638-648.

Lorenz, E., Uphoff, D., Reid, T.R., and Shelton, E. 1951. Modification of irradiation injury in mice and guinea pigs by bone marrow injections. J. Natl. Cancer Inst. 12:197-201.

Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. 2007. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J.

Majolino, I., Pearce, R., Taghipour, G., and Goldstone, A.H. 1997. Peripheral-blood stem-cell transplantation versus autologous bone marrow transplantation in Hodgkin’s and non-Hodgkin’s lymphomas: A new matched-pair analysis of the European Group for Blood and Marrow Transplantation Registry data. J. Clin. Oncol. 15:509517.

Roederer, M. 2001. Spectral compensation for flow cytometry: Visualization artifacts, limitations, and caveats. Cytometry 45:194-205.

Matsuzaki, Y., Kinjo, K., Mulligan, R.C., and Okano, H. 2004. Unexpectedly efficient homing capacity of purified murine hematopoietic stem cells. Immunity 20:87-93. Miller, C.L. and Eaves, C.J. 1997. Expansion in vitro of adult murine hematopoietic stem cells with transplantable lympho-myeloid reconstituting ability. Proc. Natl. Acad. Sci. U.S.A. 94:13648-13653.

Rossi, D.J., Bryder, D., Zahn, J.M., Ahlenius, H., Sonu, R., Wagers, A.J., and Weissman, I.L. 2005. Cell intrinsic alterations underlie hematopoietic stem cell aging. Proc. Natl. Acad. Sci. U.S.A. 102:9194-9199. Sato, T., Laver, J.H., and Ogawa, M. 1999. Reversible expression of CD34 by murine hematopoietic stem cells. Blood 94:2548-2554. Schofield, R. and Dexter, T.M. 1985. Studies on the self-renewal ability of CFU-S which have been serially transferred in long-term culture or in vivo. Leuk. Res. 9:305-313.

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Spangrude, G.J., Heimfeld, S., and Weissman, I.L. 1988. Purification and characterization of mouse hematopoietic stem cells. Science 241:58-62. Sudo, K., Ema, H., Morita, Y., and Nakauchi, H. 2000. Age-associated characteristics of murine hematopoietic stem cells. J. Exp. Med. 192:1273-1280. Szilvassy, S.J., Lansdorp, P.M., Humphries, R.K., Eaves, A.C., and Eaves, C.J. 1989. Isolation in a single step of a highly enriched murine hematopoietic stem cell population with competitive long-term repopulating ability. Blood 74:930-939. Szilvassy, S.J., Humphries, R.K., Lansdorp, P.M., Eaves, A.C., and Eaves, C.J. 1990. Quantitative assay for totipotent reconstituting hematopoietic stem cells by a competitive repopulation strategy. Proc. Natl. Acad. Sci. U.S.A. 87:87368740. Szilvassy, S.J., Ragland, P.L., Miller, C.L., and Eaves, C.J. 2003. The marrow homing efficiency of murine hematopoietic stem cells remains constant during ontogeny. Exp. Hematol. 31:331338. Tajima, F., Deguchi, T., Laver, J.H., Zeng, H., and Ogawa, M. 2001. Reciprocal expression of CD38 and CD34 by adult murine hematopoietic stem cells. Blood 97:2618-2624. Till, J.E. and McCulloch, E.A. 1961. A direct measurement of the radiation sensitivity of normal mouse bone marrow cells. Radiat. Res. 14:213222. Trevisan, M. and Iscove, N. 1995. Phenotypic analysis of murine long-term hemotopoietic reconstituting cells quantitated competitively in vivo and comparsion with more advanced colonyforming progeny. J. Exp. Med. 181:93-103. Trevisan, M., Yan, X.Q., and Iscove, N.N. 1996. Cycle initiation and colony formation in culture by murine marrow cells with long-term reconstituting potential in vivo. Blood. 88:4149-4158. Uchida, N., Dykstra, B., Lyons, K.J., Leung, F.Y.K., and Eaves, C.J. 2003. Different in vivo repopulating activities of purified hematopoietic stem cells before and after being stimulated to divide

in vitro with the same kinetics. Exp. Hematol. 31:1338-1347. Uchida, N., Dykstra, B., Lyons, K., Leung, F., Kristiansen, M., and Eaves, C. 2004. ABC transporter activities of murine hematopoietic stem cells vary according to their developmental and activation status. Blood 103:4487-4495. Wagers, A.J. and Weissman, I.L. 2006. Differential expression of alpha2 integrin separates long-term and short-term reconstituting Lin-/loThy1.1(lo)c-kit+ Sca-1+ hematopoietic stem cells. Stem Cells 24:1087-1094. Wagers, A.J., Sherwood, R.I., Christensen, J.L., and Weissman, I.L. 2002. Little evidence for developmental plasticity of adult hematopoietic stem cells. Science 297:2256-2259. Wiesmann, A., Phillips, R.L., Mojica, M., Pierce, L.J., Searles, A.E., Spangrude, G.J., and Lemischka, I. 2000. Expression of CD27 on murine hematopoietic stem and progenitor cells. Immunity 12:193-199. Wolf, N.S., Kone, A., Priestley, G.V., and Bartelmez, S.H. 1993. In vivo and in vitro characterization of long-term repopulating primitive hematopoietic cells isolated by sequential Hoechst 33342-rhodamine123 FACS selection. Exp. Hematol. 21:614-622. Yilmaz, O.H., Kiel, M.J., and Morrison, S.J. 2006. SLAM family markers are conserved among hematopoietic stem cells from old and reconstituted mice and markedly increase their purity. Blood 107:924-930. Zhang, C.C. and Lodish, H.F. 2005. Murine hematopoietic stem cells change their surface phenotype during ex vivo expansion. Blood 105:4314-4320. Zhou, S., Scheutz, J.D., Bunting, K.D., Colapietro, A.-M., Sampath, J., Morris, J.J., Lagutina, I., Grosveld, G.C., Osawa, M., Nakauchi, H., and Sorrentino, B.P. 2001. The ABC transporter Bcrp1/ABCG2 is expressed in a wide variety of stem cells and is a molecular determinant of the side-population phenotype. Nat. Med. 7:10281034.

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Analysis of the Hematopoietic Stem Cell Niche

UNIT 2A.5

Cristina Lo Celso,1, 2 Rachael J. Klein,1, 2 and David T. Scadden1, 2 1 2

Massachusetts General Hospital,Boston, Massachusetts Harvard University,Cambridge, Massachusetts

ABSTRACT Hematopoietic stem cells (HSCs) continuously replenish all blood cell lineages not only to maintain the normal rapid turnover of differentiated cells but also to respond to injury and stress. Cell-extrinsic mechanisms are critical determinants of the fine balance between HSC self-renewal and differentiation. The bone marrow microenvironment has emerged as a new area of intense study to identify which of its many components constitute the HSC niche and regulate HSC fate. While HSCs have been isolated, characterized and used in clinical practice for many years thanks to the development of very specific assays and technology (i.e., bone marrow transplants and fluorescence activated cell sorting), study of the HSC niche has evolved by combining experimental designs developed in different fields. In this unit we describe a collection of protocols spanning a wide range of techniques that can help every researcher tackling questions regarding the nature of C 2007 by John Wiley the HSC niche. Curr. Protoc. Stem Cell Biol. 3:2A.5.1-2A.5.31.
& Sons, Inc. Keywords: HSC niche r histology r intravital microscopy r homing r co-culture

INTRODUCTION Adult stem cells are responsible for the physiological maintenance of tissues and allow rapid reconstitution following injury. Stem cell fate regulation, the choice between quiescence, self-renewal, and differentiation is extremely complex and likely the result of the combination of cell-intrinsic and cell-extrinsic mechanisms. Adult tissue stem cells reside in specific locations, or niches, where neighboring cells and extracellular components not only provide a protected environment but also modulate their function (Schofield, 1978; Scadden, 2006). Hematopoietic stem cells (HSCs) give rise to all blood cells, generating progeny that follow well-defined lineage specification (Shizuru et al., 2005). Adult human and rodent HSCs reside in the bone marrow, within complex niches (Adams and Scadden, 2006). The first studies on the HSC niche were conducted with traditional HSC methodology (e.g., bone marrow transplants), when researchers noticed that alterations of the bone marrow microenvironment, and in particular of bone-lining osteoblasts, had an effect on HSC number and performance (Calvi et al., 2003; Zhang et al., 2003). The study of the HSC niche has benefited greatly from advances in the fields of hematology and general pathology, as well as from the constant advent of new technologies allowing the researcher to directly analyze tissues at the single-cell level. The protocols in this unit aim to provide basic methods for the study of HSC interactions within the bone marrow microenvironment in vivo and in vitro. Bone marrow transplants are the gold standard used to investigate the ability of HSC to self-renew and differentiate. Traditionally, whole bone marrow (WBM) and HSC have been purified from wild-type and mutant mice and transplanted into wild-type recipients. When investigating a cell-extrinsic mechanism of HSC regulation it is necessary to perform the reverse transplant, namely to use only wild-type WBM or HSC and compare their performance when injected into wild-type versus mutant recipient mice. Bone Current Protocols in Stem Cell Biology 2A.5.1-2A.5.31 Published online November 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a05s3 C 2007 John Wiley & Sons, Inc. Copyright


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marrow transplant protocols can be found in Current Protocols in Immunology. Rather than describing well-established hematopoietic protocols, we detail those more recently developed to directly analyze the HSC-niche interaction, traditionally less familiar to the hematopoietic field. Bone histological analysis can be more challenging than for other tissues but can visualize potentially any HSC antigen and bone marrow component. Basic Protocol 1 explains bone processing for the production of paraffin sections. Alternate Protocols 1 and 2 indicate the steps that differ from Basic Protocol 1 to prepare frozen sections, either from fixed, decalcified or fresh frozen bone. Basic Protocol 2 covers the basic steps of immunofluorescence on frozen sections. Small variations necessary when higher levels of signal amplifications need to be achieved are described in Alternate Protocols 3 and 4. Also immunohistochemistry methods (Alternate Protocol 5) and additional steps required when working with paraffin-embedded sections (Alternate Protocol 6) are described. In vivo imaging allows the observation of single cells within three dimensions and overcomes the limitation of the snapshot-type information if cells are monitored over time. Support Protocols 1 through 3 and 5 and 6 indicate how to prepare mice for calvarium bone marrow imaging and let them recover from the surgery. Support Protocol 4 describes live-cell staining and Basic Protocol 3 describes live-cell imaging in bone marrow. Homing and lodging assays (Basic Protocols 4 and 5) assess the ability of HSCs to engage the bone marrow niches, a critical step for the establishment of adult hematopoiesis in the bone marrow during development and for the successful outcome of bone marrow transplants. Support Protocol 4 and Support Protocol 7 indicate how to fluorescently label HSC for detection in vivo and in homing and lodging assays. Colony formation assays and HSC culture on stroma (Basic Protocol 6) are in vitro surrogates allowing some basic understanding of the influence of the microenvironment on HSC fate. NOTE: The description of HSC purification goes beyond the scope of this unit. Most of the protocols described can be performed using the hematopoietic cells of choice and not necessarily highly purified HSC. It is important to consider that cell number is a limiting factor for most assays. Lineage-negative cells are a very heterogeneous population, containing mostly nonlong-term repopulating stem cells. Very small numbers of cells are obtained when sorting lineage-negative, c-Kit-positive, and Sca1-positive cells (often referred to as LKS cells) or further HSC-enriched subpopulations of LKS cells (for a review see Bryder et al., 2006). The best cell population for each experiment must be chosen according to the variables and the aim of the experiment itself. NOTE: Most protocols described in this unit can be easily adapted to study not only the bone marrow niche but also spleen, fetal liver, thymus, and any other sites where hematopoiesis may occur; moreover, not only HSC but many other hematopoietic cell types need to reside in a specific microenvironment (or niche) to exert their function. NOTE: Most procedures described in this unit require at some stage the use of live animals. All protocols used must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations for the care and use of laboratory animals.

HISTOLOGICAL ANALYSIS

Analysis of the HSC Niche

Histological analysis, immunohistochemistry, and immunofluorescence have been routinely used to study most tissues. Hematopoietic cell biology studies have largely been performed without these techniques, due in part to the technical difficulty of sectioning through bones. Fluorescence-activated cell sorter (FACS) and cytospins were a powerful alternative to analyze bone marrow cells at the single-cell level, but they failed to give insight into characteristics of the niche because they did not allow for analysis of

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cells in their endogenous environment. The recent development of high-profile disposable blades for cryostat sectioning has made bone cryosectioning and immunostaining of bone widely accessible. In addition, use of three SLAM (signaling lymphocyte activation molecule) proteins as alternative HSC surface markers allowed a reduction in the number of antibodies (and fluorophores) needed to identify HSCs and therefore, for the first time, allowed visualization of HSC on bone marrow and spleen sections (Kiel et al., 2005).

Mouse Bone Processing for Histology: Paraffin-Embedded Blocks and Sections Paraffin-embedded blocks have traditionally been analyzed by all hospital histopathology laboratories. Relatively simple to prepare and convenient to store, paraffin blocks preserve the morphology of the analyzed tissues in very good detail.

BASIC PROTOCOL 1

Materials Mouse 3% (w/v) paraformaldehyde (PFA, see recipe) Phosphate-buffered saline (PBS, e.g., Cellgro) PBS/10% (w/v) EDTA, pH 7.5 (EDTA can be purchased from Sigma) 70% ethanol Dissecting tools: Scissors Tweezers Scalpel 24-well plate Screw-lid 15-ml conical tubes (Falcon) or 10-ml flat bottom tubes 1. Dissect the bones of interest out of the mouse, cleaning the muscle away as much as possible (for details see Lo Celso and Scadden, 2007). Whenever possible, dissect one extra ‘sentinel’ bone (see step 4). One long bone such as a tibia or femur will provide between 20 and 40 sections, so the number of bones dissected from each mouse depends on the number of sections needed. The most commonly used bone for histology is the femur, but any long or flat bone can be processed for histological analysis.

2. Fix overnight in 3% PFA at 4◦ C. The volume of PFA used is not critical. When dissecting bones from various mice, it can be useful to use 24-well pates and put up to 4 bones per well. PFA fixation works by creating protein-protein and protein-nucleic acid cross-links via methylene bridges (-CH2 -).

3. Wash once with PBS. 4. Decalcify in PBS/EDTA at 4◦ C from overnight to a few days Check that the bone is soft by bending it with tweezers (it is advisable to have an extra ‘sentinel’ bone to avoid bending the test samples and distorting the bone marrow structure inside). If the bone is not soft yet, replace the PBS/EDTA and put back at 4◦ C overnight. The length of this step depends on the bone: the calvarium is very thin and usually becomes soft in 1 to 2 days, but long bones from larger mice could take longer.

5. Wash once with PBS. 6. Put the bones in 70% ethanol in a screw-lid tube (samples can be kept like this for indefinite time) and proceed to paraffin embedding, sectioning, and hematoxylin/eosin (H&E) staining. Histology cores in most hospitals/universities are usually equipped to perform these steps with embedding machines and microtomes, and no particularly specialized equipment is necessary for bone embedding and sectioning. For a description of the instrumentation

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see Carson (1997). H&E staining can be performed also on the bench; for details see Filipe and Lake (1990). H&E staining of bone sections does not allow identification and localization of hematopoietic progenitors, but it is a first step in the analysis of bone and bone marrow morphology and it can be used, for example, to evaluate the quality of trabecular bone and the number of bone marrow blood vessels. Bone-lining osteoblasts can be easily recognized as flat to oval-shaped cells adjacent to bone and pronounced alterations in this cell type can be readily identified. ALTERNATE PROTOCOL 1

Mouse Bone Processing for Histology: Fixed Decalcified Frozen Blocks and Sections Frozen blocks require storage space at −80◦ C and are harder to section than paraffinembedded blocks, but the sections obtained are usually stained more readily since they do not require extra steps for the elimination of paraffin. Any cryostat can be used for decalcified bone sectioning, and the use of a high-profile blade is recommended to ease the procedure.

Materials PBS/20% (w/v) sucrose OCT compound (Tissue-Tek) Dry ice Cryomolds, 25 × 20 × 5–mm or 10 × 10 × 5–mm (Tissue-Tek) Cryostat (e.g., Leica, CM 3050S) Disposable high-profile blades (CL Sturkey, D554DD) Microscope slides (e.g., colorfrost plus from Fisher scientific) Paintbrush (optional) Cryogene tape system (optional, Instrumedics) Additional reagents and equipment to obtain mouse bone for processing (Basic Protocol 1) 1. Perform steps 1 to 5 of Basic Protocol 1 to obtain the mouse bone for processing. 2. Incubate in PBS/20% sucrose from a few hours to overnight at 4◦ C. This is to equilibrate the sample to a more viscous medium and to avoid swelling and breaking of BM cells while freezing.

3. Embed decalcified bone in OCT in a cryomold and freeze on dry ice. It is better to have a fair amount of OCT around the bone; typically femurs, tibias, and hips are embedded in 25 × 20–mm cryomolds, calvarium fragments in 10 × 10–mm cryomolds.

4. Store at −80◦ C until ready to cut. 5. Cut sections with a regular cryostat with a disposable high-profile blade. Mount on microscope slides. Bone and bone marrow tend to detach from each other while cutting and the best way to avoid this is to position the block so that the bone length is perpendicular to the blade. A paintbrush can be used to prevent the section from curling up while cutting (make sure not to pull it or else it will break) or use the cryogene tape system (see below). The most commonly used thickness for sections is 10 µm. Thicker sections may be cut, but a confocal microscope is necessary to analyze them properly once immunostained.

6. Store the sections at −80◦ C until ready to stain. For long-term storage it is better to keep frozen blocks rather than sections. They occupy less space and the tissue is better preserved. Analysis of the HSC Niche

Frozen sections can be stained with hematoxylin and eosin in the same way as paraffin sections. The morphology is usually better preserved by paraffin processing.

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Mouse Bone Marrow Processing for Histology: Fresh Frozen Blocks and Sections Fresh frozen blocks are the preferred starting point for immunofluorescence studies of most tissues, since they allow preservation of completely unaltered specimens. Cutting nondecalcified bones can be challenging though and should only be tried after a certain amount of experience with the cryostat has been acquired.

ALTERNATE PROTOCOL 2

Materials Mouse PBS/20% (w/v) sucrose (optional) OCT (Tissue-Tek) Dry ice Dissecting instruments Cryostat Cryostat disposable blades (e.g., D554XD Extremus from CL Sturkey) Cryogene tape system (Instrumedics) including tape and UV flashlight Cryomolds 25 × 20 × 5–mm or 10 × 10 × 5–mm (Tissue-Tek) Cryogene-coated microscope slides (Instrumedics) 1. Dissect bones out of the mouse, cleaning the muscle away as much as possible. 2. Optional: Incubate in PBS/20% sucrose from a few hours to overnight at 4◦ C. 3. Embed in OCT and freeze on dry ice (see details in Alternate Protocol 1). 4. Store at −80◦ C until ready to cut.

Figure 2A.5.1 Schematic representation of the cryogene tape system. Fresh frozen (nonfixed and nondecalcified), OCT-embedded bones are best sectioned with the cryogene tape system integrated in a cryostat. The embedded bone is positioned on the block holder so that the bone is perpendicular to the cryostat blade and the block is trimmed until the bone marrow cavity is exposed. The cryogene tape is positioned on the block so that the black line marks the bottom end of the block and touches the cryostat blade first.

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5. Cut sections with a regular cryostat equipped with a strong blade and the cryogene tape system following manufacturer’s instructions with the following modifications: a. b. c. d. e.

Open and thaw the slides for a few minutes before use. Align the black line on the tape to the bottom edge of the block (Fig. 2A.5.1). Use a roller to stick the tape on the block. Use the same roller to stick the section on the slide while positioning it. UV flash the slide twice instead of once, with a few minutes interval in between.

6. Store the sections at −80◦ C until ready to stain.

STAINING PROTOCOLS FOR TISSUE SECTIONS Immunofluorescence staining analyzed with confocal microscopy allows the best evaluation of antigen expression levels, while immunohistochemistry allows for a better analysis of tissue morphology and antigen localization although it often requires fine tuning to reduce background staining. The following protocols are general guidelines that can be followed when staining with most antibodies, but each antibody will require some optimization. Antibodies from different manufacturers differ in specificity and fluorophore intensity. Staining multiple sections at differing concentrations (for example from 1:50 to 1:1000 antibody dilution) will determine optimal staining conditions. Additionally, adjustment of the incubation times, as well as the stringency of washes, and the choice of the most appropriate blocking reagent will depend on the antibody used and the antigen of study. It is possible, in principle, to immunostain sections for any antigen. The best way to get started is to search the literature for availability of the antibody and examples of staining for the same antigen in either the same, or different tissues. Information on bone section staining is limited in the literature, and there is no single ideal protocol. For example, Jagged1 and osteopontin staining can be found in Calvi et al. (2003), and vascular staining was performed by Kiel et al. (2005). In the first case paraffin-embedded sections were stained, in the second case fresh frozen sections were stained. It is up to each researcher to troubleshoot and optimize the conditions for each new staining procedure, and it is important that the appropriate positive and negative controls (i.e., sections from other tissues already known to express the antigen and extra sections unstained or stained with secondary antibodies/streptavidin only) are prepared and carefully examined in order to trust the results obtained. A more detailed discussion on immunofluorescence and immunohistochemistry techniques can be found in Johnstone and Turner (1997). So which type of staining should be performed on which kind of sections? There is no yes or no answer. The two most common ways to prepare tissue for histological analysis are freezing and paraffin embedding. Choosing the best option depends on what components of the bone one would like to analyze and on the resources readily available. Paraffin embedding is generally used if the researcher is interested in the morphology, and/or cellularity of the tissue. Paraffin-embedded tissue is cut with a microtome and preferentially used for immunohistochemistry. Preparation of tissue for paraffin embedding includes the process of fixation and decalcification and dehydration before embedding. These steps increase autofluorescence and alter the original state of the antigens, therefore making paraffin sections less than ideal for immunofluorescence staining, and requiring antigen retrieval.

Analysis of the HSC Niche

Frozen sections are the preferred method of embedding when performing immunofluorescence staining. Fresh, frozen tissue is superior to any other embedded tissue in terms of its antigen sensitivity because the tissue is as close as possible to its original state, making the antigen detection easier and therefore not requiring antigen retrieval. However, fresh

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frozen bone is the most difficult to section because the bone is brittle and flakes easily. An easier alternative is to use fixed, decalcified, frozen sections. These are easier to section than nondecalcified bone, and they have not been dehydrated in order to be placed in paraffin, therefore antigen retrieval is not required to stain them. Because the tissue is frozen and needs to be kept at −20◦ C while sectioning, a cryostat must be employed when cutting any frozen tissue into sections, and the cryogene tape system is additionally necessary to cut fresh frozen blocks. Finally, an epifluorescence or confocal microscope is required to detect immunofluorescence. NOTE: The term “primary antibody” is used to refer to antibodies against the antigen of interest. Primary antibodies can be obtained by immunizing animals from various species, including mouse, rabbit, goat, etc. “Secondary antibodies” recognize species-specific antibodies and are therefore used to bind the primary antibody and amplify their signal.

Tissue Section Staining: Immunofluorescence on Frozen Sections Many antibodies widely used in the HSC field have been developed for flow cytometric applications and are directly conjugated to fluorophores or to biotin. Most fluorophores used for flow cytometry are easily photobleached, therefore these are not recommended for immunostaining. Biotin-conjugated primary antibodies give a slightly higher level of signal amplification than primary unconjugated purified antibodies. The use of primary antibodies raised in different species (e.g., mouse and rabbit, or rabbit and goat) coupled with species-specific secondary antibodies allows detection of two (or even more) antigens in the same section, useful for co-localization studies.

BASIC PROTOCOL 2

Materials Cryostat-cut tissue sections Phosphate-buffered saline (PBS, e.g., Cellgro) Blocking solution: PBS/10% (v/v) FBS (Invitrogen) Primary antibody of interest, purified or biotin conjugated Washing solution: PBS/0.1% or 0.2% (v/v) Tween (Sigma) Species-specific fluorophore-conjugated secondary antibody or fluorophore-conjugated streptavidin (Invitrogen, Alexa conjugates recommended) Milli-Q purified water Vectashield: either hard set (which hardens gluing the coverslip in position) or original compound (which remains liquid) with DAPI (Vector Labs) or other mounting medium Pap-pen (Fisher) Dark incubation chamber, humidified Coplin jars or beakers Thin forceps Coverslips (Fisher) NOTE: Everything is done on the bench unless otherwise stated. Incubations are in the dark in a humidified chamber, to reduce fluorophore damage and evaporation.

Prepare sections 1. Dry frozen sections on the bench for at least 30 min. Thaw one extra section to stain with only secondary antibody as negative control. Whenever possible, thaw a section from a tissue known to express the antigen analyzed as a positive control. This step is to slowly equilibrate the tissue to room temperature and to enhance section’s adhesion to the slide. This section known to express the antigen will be stained exactly like the test sections and will serve as positive control. Current Protocols in Stem Cell Biology

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2. Draw a circle around the block contour (all around the OCT/gelatin) with the pap-pen. This produces a hydrophobic membrane around the tissue and avoids solution dispersion during incubation.

3. Add enough PBS to hydrate the section, 100 to 500 µl depending on the size of the section. 4. Block with enough PBS/10% FBS to cover the section. This reduces primary antibody nonspecific binding. FBS is not the only blocking reagent one can use. Serum from various species, fish skin gelatin, and milk are widely used alternatives and generally as good as FBS. The main rule is to never use serum from the same animal that produced any of the antibodies used for the staining.

Stain sections with primary antibody 5. Apply primary antibody diluted in blocking reagent for 1 hr or alternatively at 4◦ C overnight in a humidified incubation chamber. Even though most antibodies work well diluted 1:100, dilutions between 1:50 and 1:1000 should be tried, especially when using an antibody for the first time. Usually 100 µl of antibody dilution are sufficient to cover the section. Mouse monoclonal antibodies tend to give high background on mouse sections, and it is better to use a specific Ig block first, followed by isotype-specific secondary antibody, or the Mouse on Mouse kit from Vector Labs following their instructions exactly.

6. Wash with enough PBS or PBS/0.1% or 0.2% Tween (more stringent) to cover the section three times, 5 min each.

Stain sections with secondary antibody 7. Apply secondary antibody conjugated to a fluorophore or streptavidin conjugated to a fluorophore diluted in blocking solution for 30 min. Alexa-conjugated secondary antibodies should be diluted 1:1000. Alexa-conjugated streptavidins should be diluted 1:500. Avoid Alexa 488 and any green conjugate—it will be hard to distinguish from bone marrow autofluorescence, especially if the appearance of the staining is not known in advance.

8. Wash with enough PBS or PBS/0.1% or 0.2% Tween (more stringent) to cover the section three times, 5 min each. 9. Remove the PBS and dip the slides three times in a beaker/Coplin jar containing ∼200 ml distilled water.

Mount and examine stained sections 10. Dry excess water by blotting the edge of the slide against a paper towel, being careful not to touch the sectioned tissue. 11. Drop one or two drops of Vectashield on the section; using the thin forceps slowly slide the coverslip on top of the section avoiding bubbles formation. The DAPI (4 ,6-diamidino-2-phenylindole) in the Vectashield is a nuclear counterstain that appears blue when excited with ultraviolet (UV) light.

12. Allow mounted sections to set in the dark for a few hours before observing them with the microscope. Analysis of the HSC Niche

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Tissue Section Staining: Signal Amplification Using Biotin-Conjugated Secondary Antibody

ALTERNATE PROTOCOL 3

Fixation and decalcification often cause antigen deterioration so that antibodies bind them less efficiently. Moreover, in most cases stem cells express antigens at very low levels. It is possible to increase the signal from primary antibodies by adding amplification steps. One or more molecules of a secondary antibody bind each primary antibody; in the protocol described here the secondary antibody is conjugated to numerous biotin molecules; in turn each biotin molecule is bound by streptavidin carrying the fluorophore of choice. As a consequence, more molecules of fluorophore will accumulate in proximity of the antigen than if they were directly conjugated to the secondary antibody, thus amplifying the signal.

Additional Materials (also see Basic Protocol 2) Prepared tissue sections (Basic Protocol 2) Purified unconjugated primary antibody Biotin-conjugated species-specific secondary antibody (Dako) Fluorophore-conjugated streptavidin (Invitrogen, Alexa-conjugated recommended) 1. Prepare the slides and expose sections to purified unconjugated primary antibody (Basic Protocol 2, steps 1 to 5). 2. Wash with 100 to 500 µl PBS or PBS/0.2% Tween (more stringent) three times, 5 min each. 3. Apply 100 µl/section of biotin-conjugated secondary antibody diluted 1:750 in blocking solution. Incubate for 30 min at room temperature. 4. Wash with 100 to 500 µl PBS or PBS/0.1% or 0.2% Tween (more stringent) three times, 5 min each. 5. Apply 100 µl/section of Alexa-conjugated streptavidin diluted 1:500 in blocking solution. Incubate for 30 min at room temperature. 6. Wash with 100 to 500 µl PBS or PBS/0.1% or 0.2% Tween (more stringent) three times, 5 min each. 7. Follow Basic Protocol 2, steps 9 to 12 to mount the coverslip on the slide and view the sections.

Tissue Section Staining: Amplification Using Tyramide For antigens expressed at low levels (and especially if working with fixed and decalcified tissue) it may be necessary to amplify the signal even further. Tyramide is an excellent reagent for this purpose. (Fig. 2A.5.2 represents a scheme of the various amplification strategies one can opt for.) Tyramide binds horseradish peroxidase (HRP), and it is sold conjugated to either a fluorophore or biotin. It is sold in kits from various suppliers (e.g., Perkin Elmer), usually containing specific blocking and washing solutions. It is recommended to follow the manufacturer’s instructions carefully for best results.

ALTERNATE PROTOCOL 4

Additional Materials (also see Basic Protocol 3) An HRP-conjugated reagent, either the secondary antibody or the streptavidin, both available from DAKO 3% (v/v) hydrogen peroxide Additional reagents and equipment for preparing and rehydrating sections using PBS (Basic Protocol 2)

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Figure 2A.5.2 Alternative strategies for amplification of immunofluorescence (IF) staining signal. Different levels of amplification can be used in multilayered IF in order to visualize a certain antigen (Ag) on tissue sections. The primary antibody (I Ab) recognizes the antigen and constitutes the first layer. Species-specific secondary antibody (II Ab) is the second layer. (A) II Ab used is directly conjugated to a fluorophore (II Ab Fluo, green symbol). (B) II Ab used is biotin-conjugated (gray symbol) and subsequently bound by fluorophore-conjugated streptavidin (SA, light gray symbol, Fluo, green symbol). (C) II Ab used is HRP conjugated (red symbol) and subsequently bound by fluorophore-conjugated tyramide (Tyr, blue symbol). (D) Biotin-conjugated II Ab is bound by streptavidin-conjugated HRP and subsequently by fluorophore-conjugated tyramide. (E) Biotinconjugated tyramide is used, followed by fluorophore-conjugated streptavidin. Spaces between symbols indicate reagents added in subsequent incubations, contact between symbols indicates commercially available covalently bound reagents.

1. Follow Basic Protocol 2, steps 1 to 3, to prepare and rehydrate the sections using PBS or recommended buffer. 2. Incubate the sections with 3% H2 O2 for 10 min at room temperature in the dark. Because HRP is introduced in the system, it is important to exhaust endogenous tissue peroxidase activity.

3. Wash bubbles away with PBS or recommended buffer. 4. Follow manufacturer’s instructions exactly for blocking and primary and secondary antibodies incubation steps. Select among the various levels of amplifications (see Fig. 2A.5.2) and carry out the appropriate incubations and washes. Possible amplifications include: (1) primary antibody, then HRP-conjugated secondary antibody, then Tyr-fluorophore conjugated reagent; (2) Primary antibody, then biotinconjugated secondary antibody, then streptavidin-conjugated HRP, then Tyr-fluorophore conjugated reagent; (3) Primary antibody, then biotin-conjugated secondary antibody, then streptavidin-conjugated HRP, then Tyr-biotin, then Alexa-conjugated streptavidin. The more amplification that is used, the more non-specific background staining may be produced. It is critical to have a negative control slide stained with a matching isotype primary antibody or with secondary antibody and reagents only. Analysis of the HSC Niche

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Tissue Section Staining: Immunohistochemistry Immunohistochemistry works the same way as immunofluorescence, but it terminates with a colorimetric reaction in which an enzyme converts a soluble substrate into an insoluble colored precipitate. One enzyme typically used for this reaction is horseradish peroxidase (HRP), introduced in the system by using a secondary antibody conjugated to HRP or a secondary antibody conjugated to biotin followed by streptavidin-HRP. Alternatively, alkaline phosphatase (AP) can be used, and in this protocol every time HRP is mentioned it can be replaced with AP. Typical substrates for HRP are DAB (3,3diaminobenzidine) and AEC [3-amino, 9-ethyl-carbazole, red precipitate (Calvi et al., 2003)]. NBT/BCIP solutions are used with AP to generate a blue precipitate [for more details see Filipe and Lake (1990) and Lo Celso et al. (2004)].

ALTERNATE PROTOCOL 5

Materials Frozen sections Phosphate-buffered saline (PBS, e.g., Cellgro) 3% (v/v) H2 O2 Blocking solution: PBS/10% (v/v) FBS Primary antibody of interest, purified or biotin conjugated Washing solution: PBS/0.1% or 0.2% (v/v) Tween Secondary antibody, HRP- or biotin-conjugated (Dako) Streptavidin, HRP-conjugated (DAKO), optional DAB (see recipe) Milli-Q purified water Hematoxylin Pap-pen Beakers or containers holding at least 200 ml distilled water for final washes Prepare samples 1. Dry frozen sections on the bench for at least 30 min. Everything is done on the bench unless stated otherwise.

2. Draw a circle around the section with the pap-pen. 3. Add enough PBS to hydrate the section, 100 to 500 µl depending on the size of the section.

Block endogenous HRP activity 4. Block endogenous HRP activity with 100 to 500 µl 3% H2 O2 for 10 min. 5. Wash bubbles away with 100 to 500 µl PBS. Repeat the wash if bubbles persist. 6. Block with 100 to 500 µl PBS/10% FBS for 1 hr.

Expose to primary antibody 7. Apply primary antibody diluted in PBS/10% FBS, 100 µl/section for 1 hr. 8. Wash with 100 to 500 µl PBS or PBS/0.1% or 0.2% Tween (more stringent) three times, 5 min each.

Expose to secondary antibody 9. Apply HRP- or biotin-conjugated secondary antibody diluted in blocking solution, 100 µl/section for 30 min. For dilutions it is best to check the manufacturer’s instructions. If the primary antibody is biotin conjugated, skip this step and go to step 11.

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10. Wash with PBS or PBS/0.1% or 0.2% Tween (more stringent) three times, 5 min each. 11. Apply HRP-conjugated streptavidin diluted 1:100 in blocking solution or according to manufacturer’s instructions. Incubate using 100 µl/section for 30 min. 12. Wash with 100 to 500 µl PBS or PBS/0.1% or 0.2% Tween (more stringent) three times 5 min each.

Perform detection reaction 13. Apply 100 to 500 µl DAB reaction solution. The longer the DAB is left on the slide the more brown product forms. This enhances the staining but also increases the background and if left for too long the whole section will turn brown. Standard reaction times are from a few seconds to 5 min. It is good to check how the reaction is proceeding by putting the slide onto something white every now and then.

14. Stop DAB reaction by removing DAB solution and adding a 100- to 500-µl drop of distilled water. Wash sections three times by immersing them in water (water can be in a beaker or other container holding at least 200 ml distilled water), 2 min each. 15. Counterstain sections. Hematoxylin is a good nuclear counterstain to show the morphology of the tissue. Histopathology cores usually do very good hematoxylin staining and mounting, but it can be done on the bench too (Filipe and Lake, 1990).

ALTERNATE PROTOCOL 6

Tissue Section Staining: Immunofluorescence and Immunohistochemistry on Paraffin Sections Generally paraffin sections tend to have higher autofluorescence than frozen sections and are more suitable for immunohistochemistry, but they can also be used for immunofluorescence. Some antibodies will recognize the antigen on paraffin sections as efficiently as on frozen sections, but frequently the antigen retrieval step is required right after deparaffinizing and before starting the staining. There are many methods and they can be better or worse depending on the antibody. They are mostly based on boiling the slides for a while in a buffer (the most commonly used is citrate buffer, here described in detail) and cooling them down slowly afterwards. Here we describe citrate buffer antigen retrieval, but many alternative protocols can be found online on the Information Center for Immunohistochemistry Web site (IHC World, http://www.ihcworld.com/epitope%20retrieval.htm).

Additional Materials (also see Basic Protocol 5) Xylene (Fisher) 80% and 100% ethanol (Fisher) Distilled water Na citrate or other antigen retrieval buffer (see recipe) 500-ml to 1-liter beaker or microwave-proof container with lid Microwave oven Additional reagents and equipment for tissue section staining (Alternate Protocol 5) 1. Deparaffinize the section by dipping the slide in xylene three times, 5 min each, and rehydrate the section by incubating in an ethanol series: Analysis of the HSC Niche

100% ethanol, three times, 2 min each 80% ethanol, once for 2 min

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H2 O2 twice, 2 min each PBS once for 5 min. Most histopathology cores are equipped for this step but it can be done in house too provided there is a fume hood for xylene handling and use.

2. Boil the slides in antigen retrieval buffer in a microwave-proof container for 30 sec to 30 min, depending on the antigen and the microwave power. It is critical that the sections do no get dry in the process (use a lid and plenty of buffer).

3. Once removed from the microwave, let slides cool on the bench for 30 min before washing with PBS and starting the staining. 4. Follow the general immunohistochemistry protocol (Alternate Protocol 5) from step 2 or immunofluorescence protocols (Basic Protocol 2 and Alternate Protocols 3 and 4) from step 2.

IN VIVO IMAGING When planning an in vivo imaging experiment, it is important to keep in mind that the signals produced by the fluorophores have to be very bright in order to be detected. To get a rough estimate of the level of brightness needed, cells in suspensions can be observed first. Roughly ten times extra power will be needed to detect the same kind of signal within the bone marrow or ten times brighter signal will be required to detect similar signal using the same settings in vivo.

Bone Marrow Live Imaging Calvarium bone marrow is an active site of hematopoiesis in both humans and mice and is the most accessible site for imaging because it is exposed by simply making an incision in the scalp.

BASIC PROTOCOL 3

NOTE: The term calvarium refers to all bones forming the top part of the skull.

Materials Mice injected with labeled cells (Support Protocol 4) Anesthetic of choice (Support Protocols 1 through 3) Fluorescent vascular dye (e.g., fluorescently labeled dextran or Angiosense, see recipe) Stained cells of choice, optional Phosphate-buffered saline (PBS, e.g., Cellgro) Methocell (optional, OmniVision) Buprenorphine (Buprenex) Hair clipper or sharp scissors, optional Surgery thin tweezers and scissors Suturing thread [e.g., Ethilon 6-0 (0.7 metric) PC-3, Ethicon, no. 1966G] or glue (Vetbond, 3 M) Suture needle Warmed stage or warmed cylindrical holder Appropriate intravital microscope (e.g., Olympus IV100) Anesthetize the mouse 1. Anesthetize the mouse using the method of choice (Support Protocols 1 to 3). 2. Check that the mouse is fully anesthetized by toe pinch reaction.

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Inject vascular dye 3. Optional: If observation of the vasculature is required, intravenously inject the vascular dye of choice (see Reagents and Solutions for details about vascular dye choice; both tail vein and retro-orbital injections work fine). If you are going to observe cells homing to the bone marrow or circulating, they can also be injected at this point. If it is crucial to image the cells right after injection, at this point it is possible to simply position a catheter with some heparin in the mouse tail vein (Chabner et al., 2004) and the cells can be injected after the mouse has been correctly positioned under the objective (step 10).

Perform surgery to expose calvarium 4. Shave head hair with a hair clipper or cut it with sharp scissors. Steps 5 to 8 are optional and used if imaging is not the final step in the experiment.

5. With sharp scissors make a midline incision in the scalp, from between the ears and going all the way to between the eyes and close to the nose. If the imaging session is the end of the experiment you might just cut away the necessary part of skin since you don’t need to worry about suturing afterwards.

6. Insert suturing thread through one side of the cut skin, just above the eye, from the incision side to the outside side. 7. Fold the thread under the jaw of the mouse and go through the other side of open skin, again above the eye and from the incision side to the outside side. 8. Cut needle away from thread and make a knot with the two ends under the jaw of the mouse. This will keep the incision open so that the area available for imaging is wider and unobstructed. 9. Use water or PBS to rinse away any hair accidentally sticking on the skull. Hair fragments produce considerable light scattering on the surface and shadows underneath so it is important to clean the skull very well.

10. Keep the mouse warm while it is anesthetized. Position it either onto a warmed stage or into a warmed cylindrical holder. The mouse is ready for imaging. Depending on the microscope employed for the experiment, further preparations may be necessary at this step. Most intravital microscopes are equipped with dry long-distance lenses requiring no further tissue preparation. However, even if using a long-distance objective, once the scalp has been opened the bone tends to dry, which can lead to tissue decay and impair the penetration of the laser beam through the bone. The use of a gel (e.g., Methocell) to maintain moisture is recommended, but the most appropriate reagent to use may vary depending on the microscope objective. Some intravital microscopes work with immersion lenses hence it is necessary to keep a fair amount of PBS or gel on the calvarium. It is possible to position a coverslip on the head of the mouse if water immersion objectives are used. In this case Methocell is a good gel to maintain the calvarium’s physiological moisture level. When positioning the mouse under the objective, it is helpful to try and center the central vein under the objective and to align the mouse so that the central vein will be a straight line either vertical or horizontal within the observation field. Locating the central vein helps to keep track of where the imaging is performed within the calvarium.

Image the bone marrow 11. Image the bone marrow of the calvarium. Analysis of the HSC Niche

Ideally, a two-photon microscope is used to visualize the bone by collagen second harmonic generation (e.g., exciting at 860 nm and acquiring at 420 nm) and fluorophores (e.g., for GFP exciting at 920 nm and acquiring at 460 nm). Near-infrared dyes have

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to be imaged with confocal microscopes since most EP lasers do not reach long enough wavelengths. The combination of lasers and filters used depends heavily on the microscope available. Even simple epifluorescence microscopes can produce images of high quality.

12. If the mouse has to survive the imaging, close the incision once imaging has been completed. Use stitches or 3 M Vetbond glue (see Support Protocols 5 and 6) to close the scalp. It is important to suture the incision well in order to reduce the formation of scar tissue, which makes it difficult to image repeatedly. To reduce inflammation and scarring, an antibiotic cream can be spread on the calvarium before suturing (any topical antibiotic cream sold in a pharmacy will work).

13. Monitor the animals during the post-operative period. Keep the animal warm until it wakes from anesthesia. Treat it with analgesic (e.g., 0.05 to 1 mg/kg buprenorphine HCl administered intramuscularly every 12 hr for 48 hr). It is safe to image the mouse again the following day. Multiple imaging sessions per day increase the probability of mouse death due to anesthetic overdose. The dose of ketamine/xylazine should be reduced at each repeated injection.

Mouse Anesthesia: Ketamine/Xylazine Mice must be anesthetized in order to surgically expose the calvarium and additionally restrained to stabilize their head during imaging. Irrespective of the anesthesia method chosen, the depth of anesthesia of the mouse has to be monitored and the body temperature of the mouse has to be maintained throughout the experiment (see methods below). A comparison between various methods of anesthesia can be found in Arras et al. (2001).

SUPPORT PROTOCOL 1

Ketamine/xylazine cocktail administration is a widely used anesthesia method. Dosage varies depending on the size of the mouse and on the strain. This method is relatively simple as it requires only one bottle of mixed anesthetic, which can be stored for several months at room temperature, and no specific machinery for the administration of the anesthetic.

Materials Mouse Ketamine/Xylazine cocktail (see recipe) Syringe Balance Heated pad 1. Weigh the mouse and prepare a syringe with the correct dose of anesthetic. The recommended dosage is 80 mg/kg ketamine and 12 mg/kg xylazine, so inject 50 µl of the prepared cocktail in a mouse weighing 25 grams. The amount of ketamine/xylazine cocktail to inject has to be adjusted according to mouse weight and strain.

2. Inject ketamine/xylazine intraperitoneally in the mouse. 3. Keep the mouse warm on a heated pad. 4. Check that the mouse is anesthetized by absence of reaction to toe pinch. This anesthesia lasts for ∼1 to 1.5 hr. Somatic Stem Cells

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SUPPORT PROTOCOL 2

Mouse Anesthesia: Avertin Avertin keeps the mice anesthetized for longer than the ketamine/xylazine cocktail and it is harder to overdose.

Materials Mouse Avertin solution (see recipe) Syringe Balance Heated pad 1. Weigh the mouse and prepare the syringe with the appropriate dose of avertin (0.025 ml/g body weight). 2. Inject avertin intraperitoneally. 3. Keep the mouse warm on a heated pad. 4. Check that the mouse is anesthetized by absence of reaction to toe pinch. This anesthesia lasts for ∼1 to 1.5 hr. SUPPORT PROTOCOL 3

Mouse Anesthesia: Isoflurane This is a very quick and simple inhalation anesthesia; the mouse can be kept anesthetized for a few hours and wakes up quickly once the isoflurane is removed.

Materials Mouse Isoflurane (e.g., Nicholas Piramal or Henry Shein) Oxygen (e.g., Nicholas Piramal or Henry Shein) Isoflurane anesthesia machinery (e.g., Henry Shein) 1. Place the mouse in the isoflurane anesthesia machinery. 2. Adjust the isoflurane flux to 0.8 to 1.5 liter/min and the isoflurane vaporizer to 2% to 3%. The anesthesia machinery mixes isoflurane with oxygen to obtain the desired isoflurane concentration.

3. For maintenance, adjust the flowmeter to 400 to 800 ml/min. 4. Continue anesthesia, maintaining body temperature, until the procedure is complete. SUPPORT PROTOCOL 4

Analysis of the HSC Niche

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Live Cell Staining Lipophilic dyes are currently the best option to mark cells for in vivo imaging. Invitrogen offers a full range of these dyes which absorb energy and emit signal in various regions of the light spectrum. For example, DiD (1,1 -dioctadecyl3,3,3 ,3 -tetramethylindodicarbocyanine perchlorate) is excited with red light (640 nm) and its emission peaks around 650 nm, while DiI (1,1 -dioctadecyl-3,3,3 ,3 tetramethylindocarbocyanine perchlorate) is excited with green light (530 nm) and its emission peaks around 580 nm. Some dyes have a broader excitation/emission spectrum, so make sure to pick dyes sufficiently far apart in at least one of the two spectra when planning simultaneous staining and imaging of multiple populations. Good spectra viewers allowing comparison of different dyes are available on both the Invitrogen and BD Biosciences Web sites (http://probes.invitrogen.com/resources/spectraviewer/ and http://www.bdbiosciences.com/spectra/) and the Invitrogen Web site contains detailed tutorials about excitation and emission spectra (http://probes.invitrogen. com/resources/education/tutorials/2Spectra/player.html). Current Protocols in Stem Cell Biology

Different cell types are stained optimally using different conditions and dyes. Alternative dyes widely used for cell tracking experiments include PKH dyes from Sigma (Askenasy and Farkas, 2002; Askenasy et al., 2002), Calcein-AM (Cavanagh et al., 2005), and Cell Tracker Orange (Mazo et al., 2005) from Invitrogen. The manufacturer’s instructions and the abovementioned papers contain a detailed description of staining methods for each dye. When choosing the fluorophores, it might be worth considering that near infrared fluorophores are the ones detected deepest in live tissues, therefore allowing detection of the greatest amount of tissue. When staining multiple cell types to track simultaneously it is necessary to choose dyes with different emission spectra in order to minimize overlap in signal detection. Also, when imaging cells and vasculature at the same time it is important to minimize the overlap between cellular and vascular dyes. NOTE: CFDA-SE (carboxyfluorescein diacetate, succinimidyl ester, Invitrogen) is an intracellular dye generating a signal similar to GFP. It has been used for several localization studies, but more recently toxicity issues have emerged, therefore it should not be the dye of choice, especially if comparing two or more populations stained with different dyes and injected simultaneously. NOTE: The protocol below is recommended for staining of live HSC for any kind of assay.

Materials Cells of choice Phosphate-buffered saline (PBS) DiD or DiI (Invitrogen) 37◦ C incubator 1.0-ml insulin syringe equipped with a 31-G needle 1. Resuspend cells of choice at an approximate density of 106 cells/ml in PBS with no serum. 2. Add DiD to a final dilution of 5 µM and quickly mix by pipetting/vortexing to avoid DiD coming out of solution. 3. Incubate for 10 min at 37◦ C in the dark. 4. Fill the tube with PBS and centrifuge for 5 min at 500 × g, room temperature. The pellet should be light blue.

5. Resuspend the cells in 100 to 300 µl of PBS. 6. Collect the cells in an insulin syringe (preferable to avoid losing cells in the needle dead space, especially when injecting small numbers of cells) and, without changing needle, inject them intravenously as quickly as possible by tail vein injection (retroorbital injection is advisable only if the mouse is already anesthetized for immediate imaging).

Suturing the Scalp After imaging is complete, the scalp should be carefully sutured to minimize the scarring if repeated imaging will be used.

Materials Mouse from imaging experiment Suturing thread [Ethilon 6-0 (0.7 metric) PC-3, Ethicon, no.1966G] Suture needle Tweezers 1. To suture the mouse, cut away the thread that was holding the incision open.

SUPPORT PROTOCOL 5

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2. With a new needle and thread, stitch through the two sides of open skin. 3. Cut threaded needle away, leaving enough thread on it for both the next stitch and for tying a knot. 4. Grab left thread with right tweezers. 5. Loop thread around left tweezers twice loosely (or once and repeat steps 5 to 7 twice). 6. Grab right thread with left tweezers. 7. Pull tight to stabilize the knot. 8. Make as many stitches as necessary to completely close the wound. Usually four to six stitches are sufficient to suture the scalp. SUPPORT PROTOCOL 6

Gluing the Scalp Instead of suturing the mouse it is possible to use skin glue (e.g., 3 M Vetbond). The glue is less of an irritant than stitches, but it is very important to stick the entirety of the skin sides together to avoid scaring. It is crucial to avoid pouring the liquid glue in the eyes or nostrils.

HOMING AND LODGING ASSAYS The terms homing and lodging are used almost interchangeably to refer to the localization of hematopoietic cells to the bone marrow following transplant or, physiologically, following migration from fetal liver. Homing refers generally to the ability of HSC to reach the bone marrow, while lodging is somehow one step beyond and involves the ability of HSCs to engage the niche and remain in their specialized microenvironment close to the endosteal surface (Wolf, 1974; Adams et al., 2006). Homing and lodging assays are therefore used to evaluate the ability of control and test cells to migrate to the bone marrow and the endosteal surface of wild-type and mutant recipients. Irradiation profoundly alters bone marrow anatomy and therefore it is not recommended when a close look at the localization of the injected cells is necessary. Practically, irradiation is administered to recipient mice in homing assays, where the exact localization of cells within the bone marrow is not taken in consideration, but not to recipient mice in lodging assays, since disruption of bone marrow anatomy would impair evaluation of HSC localization within the bone marrow. BASIC PROTOCOL 4

Homing Assay This assay determines the ability of the test cells to migrate to any region within the bone marrow. The same procedure could be used to analyze homing to the spleen or other organs. Usually either total bone marrow or lineage-depleted bone marrow cells are injected (for instructions on cell preparation see Adams et al., 2006 and Lo Celso and Scadden, 2007). If lineage depletion is used, it is important to check and record the efficiency of lineage depletion before each injection by staining a small aliquot of total bone marrow and depleted cells with a cocktail of antibodies against differentiated cells (Lin cocktail) and analyzing the samples by flow cytometry. Since the efficiency of lineage depletion tends to be highly variable, injecting total bone marrow and staining for lineage afterwards is likely to give the most consistent results over repeated experiments.

Materials

Analysis of the HSC Niche

Mice Stained cells of choice PBS/2% (v/v) FBS Lineage cocktail antibodies (equal amounts of Mac1, Gr1, Ter119, CD4, CD8, CD3, and B220 biotinylated antibodies, BD Pharmingen), optional Fluorophore-conjugated streptavidin (BD Pharmingen)

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Gamma irradiator Needles and syringes Dissecting instruments 40-µm filters (BD) FACS tubes (BD) FACS machine (Calibur, BD) Additional reagents and equipment for harvesting cells from the donor mice (Lo Celso and Scadden, 2007), labeling the cells appropriately (Support Protocols 4 and 7) and counting cells using a hemacytometer (Phelan, 2006) Irradiate mice 1. Lethally irradiate recipient mice (∼10 Gy for C57Bl/6 or 3.5 Gy for NOD/SCID). The type of irradiator available is strictly regulated by national/federal guidances and the radiation safety department of each hospital/university usually deals with irradiator installation. One commonly found type of irradiator uses Cesium 137 as gamma irradiation source. The irradiation kills most bone marrow cells, therefore making space for the test cells that are injected.

Prepare cells for injection 2. Prepare the cells for injection: a. Harvest them from the donor mice (Lo Celso and Scadden, 2007). b. Label the cells appropriately (see Support Protocols 4 or 7). c. Count the cells with a hemacytometer (Phelan, 2006). 3. At a time point 4 to 24 hr following irradiation, inject the labeled cells intravenously. A typical injection would be 5 × 106 total bone marrow cells and 1 × 106 lineage-depleted cells per mouse.

Sacrifice mice 4. At a time point 6 hr following injection of cells, sacrifice the recipient mice. 5. Dissect one femur from each mouse. Alternatively, dissect the organ of interest and prepare a single-cell suspension.

Collect the cells 6. Crush the femur in 5 to 10 ml PBS/2% FBS. 7. Filter the cell suspension through a 40-µm filter. 8. Centrifuge 5 min at 400 × g, room temperature, resuspend in 1 ml PBS/2% FBS.

Stain for lineage markers (optional) 9. Incubate 200 µl of cells/recipient mouse in FACS tubes with 4 µl of biotin-conjugated lineage antibodies cocktail for 30 min at 4◦ C. This step allows distinguishing between differentiated and more primitive cells homed to the bone marrow when total bone marrow is injected. The staining can be performed directly in the FACS tube.

10. Wash twice with 4 ml PBS/2% FBS. Fill the tubes, spin 5 min at 500 × g, room temperature, discard the PBS. 11. Resuspend in 200 ml PBS/2% (v/v) FBS with streptavidin conjugated to the appropriate fluorophore (e.g., a fluorophore different from the cell dye used; streptavidin is typically diluted 1:500) for another 30 min at 4◦ C.

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12. Fill each tube with 4 ml PBS/2% FBS. 13. Centrifuge 5 min at 500 × g, room temperature, discard the PBS, and resuspend the cells in 200 ml PBS/2% (v/v) FBS.

Analyze suspension 14. Analyze the cell suspension (step 8 or step 13) by flow cytometry. BASIC PROTOCOL 5

Lodgment Assay This assay is structured essentially in the same way as the homing assay, but the location where the injected cells reside within the tissue of interest is investigated in more detail using histological analysis. The recipient mice are not irradiated in order to avoid any destruction of the bone marrow structure.

Materials Cells of choice Phosphate-buffered saline (PBS; e.g., Cellgro) Vectashield Coverslips Thin forceps (to handle coverslips) Additional reagents and solutions for labeling cells (Support Protocols 4 and 7) and dissecting the femur and processing for histological analysis (Basic Protocol 1 or Alternate Protocol 1) 1. Prepare the cells of choice by dissecting the mouse, crushing the bones, and (optional) performing lineage-depletion (Lo Celso and Scadden, 2007). Make sure to have enough cells to inject at least 50% extra mice (cells will be lost at each centrifugation step). Due to limiting cell numbers, total bone marrow mononuclear cells or lin-negative fraction is most commonly used for this assay.

2. Label the cells (see Support Protocols 4 or 7) and resuspend them in PBS. 3. Inject the cells of choice intravenously (tail vein injection) into the recipients in 200 to 500 µl PBS. Typically 5 × 106 total bone marrow or 500,000 lineage-depleted cells are injected.

4. At a time point 6 hr later sacrifice the mice, dissect the femurs or the organ of choice and process for histological analysis (follow Basic Protocol 1 or Alternate Protocol 1, depending on the availability of the instruments necessary for paraffin or frozen blocks processing and sectioning). CFDA-SE and DiD remain bright following fixation and decalcification, so it is not necessary to section nonfixed, nondecalcified blocks (Alternate Protocol 2). If the tested cells are GFP-labeled, it is advisable to avoid fixation, which decreases GFP brightness. It is advisable to work with fresh frozen blocks and sections (Alternate Protocol 2) or to analyze fixed decalcified paraffin or frozen sections using an anti-GFP antibody to stain for the injected cells (e.g., use Abcam’s rabbit anti-GFP antibody # ab-290 and follow Basic Protocol 2 or Alternate Protocol 6).

5. Once the sections are prepared, hydrate them with PBS for a few minutes. 6. Mount sections with vectashield (plus DAPI) and add a coverslip. 7. Analyze the sections. Analysis of the HSC Niche

The injected cells are brightly labeled so no other staining is needed to identify them (Fig. 2A.5.3), unless they express low levels of GFP or other fluorescent proteins.

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Figure 2A.5.3 Lodging assay. Picture of a section obtained from the hip bone of a non-irradiated mouse injected with DiD stained lineage-negative cells 6 hr prior to sacrifice. Green is cytoplasmic bone and bone marrow cells’ autofluorescence signal, blue is DAPI nuclear counterstain, purple is DiD membrane dye. Arrow indicates a lineage-negative cell. Scale bar is 100 µm.

Staining of Cells with CFDA-SE The most widely used label for homing assays is CFDA-SE (Nilsson et al., 2001; Adams et al., 2006) but recently there have been indications of some degrees of toxicity for CFDA-SE, hence a different dye is recommended. However, if the spectrum of other dyes is not easily analyzable with the available resources, CDFA-SE is still a valid marker since all cells (test and control) are stained in the same way and no comparison is made between cells stained with CFDA-SE and different dyes. For more details see Support Protocol 4.

SUPPORT PROTOCOL 7

Materials Cells of choice Phosphate-buffered saline (PBS), 37◦ C 100 µM CFDA SE stock (see recipe) Medium (e.g., DMEM, Invitrogen) supplemented with 10% (v/v) FBS, 37◦ C Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) Additional reagents and equipment for preparing cells of choice (Basic Protocol 5) 1. Centrifuge cells 5 min at 500 × g, room temperature to pellet and resuspend them in 1 ml of 37◦ C PBS. 2. Add 50 µl of the 100 µM CFDA-SE stock to the resuspended cells (to make a 5 µM working dilution).

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3. Incubate the cells 15 min at 37◦ C in the dark (e.g., in a cell culture incubator). 4. Centrifuge the cells 5 min at 500 × g, room temperature and resuspend the pellet in 1 ml of prewarmed medium (e.g., DMEM/10% FBS). 5. Incubate for a further 30 min at 37◦ C. 6. Centrifuge the cells 5 min at 500 × g, room temperature and resuspend the pellet in CMF-PBS in a volume suitable for injection (typically 5 × 106 WBM or 5 × 105 lin− cells/300 CMF-PBS/mouse). The cells are ready to be used. BASIC PROTOCOL 6

HSC-STROMA CO-CULTURE: CAFC/LTC-IC ASSAY This is the most versatile in vitro assay to assess the ability of potentially any stroma cell type to support hematopoietic cell growth [cobblestone area-forming cell (CAFC) assessment] and differentiation [long-term culture-initiating cell (LTC-IC) scoring]. Not only wild-type versus mutant primary stroma cells can be compared, but also several cell lines can be used to assess the effect of a wider range of genetic modifications on the interaction between stroma and hematopoietic cells, for example AFT024 (Charbord and Moore, 2005) or M2.10B4 (Sutherland et al., 1991; Bouzianas, 2003). OP9 cells have been used in protocols differentiating ES cells into hematopoietic cells (Kitajima et al., 2006) and OP9DL1, transduced to express Delta-like 1 Notch ligand have been used to culture and differentiate T cells (Taghon et al., 2005). The term stroma is usually used to refer to the nonhematopoietic component of the bone marrow, therefore it is a highly heterogeneous cell population. Practically, stromal cells are separated from hematopoietic cells due to their ability to adhere to culture plates, a property shared by cells of bone and vascular lineages, but also macrophages. No specific protocol for the isolation of a specific stroma cell type is widely accepted yet. Human stromal cells can be plated from cord blood or bone marrow biopsies. The stroma cell line M2.10B4 can be grown using the same method as for bone marrow-derived primary stroma. CAFC/LTC-IC assay was developed as a way to assess HSC number in a given population. While limiting dilution and serial transplants are now preferentially indicated as gold standard to compare HSC numbers in control and test population, CAFC/LTC-IC assay can be re-visited and used to evaluate differential interaction between hematopoietic cells and test versus control stroma.

Materials Mice Collagenase (Worthington) PBS/2% and 20% FBS Long-term culture medium (H5100 for human or M5300 for murine cells, see recipe) Trypsin with EDTA (Cellgro # 25-053-CI) dH2 O Hematopoietic cells of choice Methylcellulose-containing medium (see recipe)

Analysis of the HSC Niche

Dissecting tools Mortar and pestle (Fisher) 70-µm filters (BD) Scissors Heated water bath with shaker

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25- or 75-cm2 flasks (Nunc) Irradiator (see Basic Protocol 4 for more details) 96-well plate 33◦ C incubator 5% CO2 L-Calc software (free from Stem Cells Technologies) Additional reagents and equipment for dissecting bones from mice (Lo Celso and Scadden, 2007) Dissect bones 1. Dissect all bones from one or more mice (see Lo Celso and Scadden, 2007). 2. Crush all bones in the mortar and obtain bone marrow by filtering trough 70-µm filters. Keep the filtered cells in the refrigerator while waiting. IMPORTANT NOTE: Steps 3 to 8 are optional, but they allow for a higher yield of osteoblasts (Zhu et al., 2007).

3. Collect all pieces of bone that remained in the mortar and cut them in smaller pieces using scissors. 4. Incubate the bone fragments with 10 ml of 3 mg/ml collagenase per mouse at 37◦ C with agitation for 30 to 45 min. 5. Fill the tube with PBS/20% FBS and centrifuge 5 min at 500 × g, room temperature. 6. Aspirate the medium and vortex the bone fragments vigorously for ∼20 sec. 7. Resuspend in PBS/2% FBS, 5 ml per mouse. 8. Filter through a 70-µm filter and pool with the mononuclear cells obtained from crushing (step 2).

Culture stromal cells 9. Culture stroma cells at 5 × 106 cells/ml in long-term culture medium in 25-cm2 (up to 6 ml) or 75-cm2 flask (up to 15 ml). 10. Let stromal cells grow in the same flasks from step 9 until confluent (human ∼4 weeks; murine ∼2 weeks) changing half of the medium every 3 to 4 days. 11. Trypsinize cells from flask with 0.25% trypsin/0.53 mM EDTA for a few minutes at 37◦ C. Add medium to trypsinized cells to stop trypsinization. 12. Centrifuge 5 min at 500 × g, room temperature and resuspend in medium at ∼106 cells/ml.

Initiate stromal cell cultures for assay 13. Irradiate the cell suspension with 15 Gy. 14. Dilute cells to 2.5 × 105 /ml in long-term medium and plate out 100 µl of cells (i.e., 2.5 × 104 ) in the middle 6 × 10 wells of 96-well plate. 15. Add 200 µl dH2 O to the remaining wells. 16. Incubate at 33◦ C (the plate can now be left for 1 to 7 days before being used).

Set up assay 17. Obtain mouse or human total bone marrow mononuclear cells or the preferred purified subpopulation of hematopoietic cells. For isolation of mouse bone marrow cells/HSC see Lo Celso and Scadden (2007); for human HSC isolation see Pearce and Bonnet (2007).

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18. Prepare five serial two-fold dilutions of cells in long-term medium. For CD34+ human cells, prepare at 10,000, 5,000, 2,500, 1,250, and 625 cells/ml each in 5100 µl medium. For murine bone marrow mononuclear cells, prepare at 2 × 106 , 1 × 106 , 5 × 105 , 2.5 × 105 and 1.25 × 105 cells/ml in each in 5300 µl medium (note: for other cell types you may need to adjust the seeding number).

19. Add 50 µl of each dilution to each of 12 wells in the 96-well plate. This means for CD34+ cells you will be adding 500, 250 etc. cells per well. For murine MNCs, 100,000, 50,000, 25,000 etc. per well. Figure 2A.5.4 is a scheme of the plate set up.

20. Incubate at 33◦ C for 5 weeks, replacing 75 µl of medium/well weekly. Also check the wells with water and replenish if necessary.

Score CAFCs 21. Score wells either positive or negative for CAFCs (wells are positive if they include a cobblestone of >5 cells). 22. Calculate CAFC frequency using L-Calc software to obtain a readout of hematopoietic cell proliferation.

Determine LTC-IC frequency 23. Centrifuge plate 5 min at 500 × g, room temperature. 24. Carefully aspirate all medium.

Analysis of the HSC Niche

Figure 2A.5.4 Layout of a CAFC/LTC-IC experiment. Water is distributed in all peripheral wells of a 96-well plate (blue circles) and serial dilutions of mouse whole bone marrow mononuclear cells (WBM MNC) or human CD34+ cells are seeded on top of stroma in the central wells. Black-to-white gradient indicates wells seeded with decreasing number of cells.

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25. Add 100 µl of methylcellulose-containing medium (either M3434 or H4435). 26. Incubate plate for 2 weeks at 37◦ C. 27. Score wells either positive or negative for LTC-ICs (wells are positive if they contain a colony with >20 cells). 28. Calculate LTC-IC frequency with L-Calc to obtain a readout of differentiation.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Antigen retrieval buffer Dissolve 0.96 g citric acid in 500 ml H2 O2 Use NaOH to adjust the pH to 6 Use within the day It is possible to buy premade 10× buffers from BioGenex. Alternatively, 10 mM citrate buffer, pH 6.0, can be prepared just before starting the staining.

Avertin Diluent recipe: 0.8% (w/v) NaCl 1 mM Tris·Cl, pH 7.4 0.25 mM EDTA Check the pH and adjust to pH 7.4 Prepare avertin stock by mixing 1 g tribromoethanol in 0.5 ml tert-amyl alcohol (2 methyl-2-butanol). Dissolve by heating to 37◦ C overnight. Store wrapped in foil (light sensitive solution; alternatively use brown glass bottle) up to 6 months at 4◦ C (decomposition can result from improper storage). The mixture should be clear, if solution becomes opaque over time, it should be warmed to dissolve any particulate.

Prepare working stock avertin (this solution should be prepared weekly) by diluting 60 µl of stock in 5 ml PBS or diluent.

Filter with 0.22-µm filter syringe Store up to 6 months at 4◦ C, in a foil-wrapped or brown bottle CFDA-SE stock Dissolve the contents of component A in 90 µl of component B (DMSO) of the Invitrogen CFDA-SE cell tracer kit (no. V12883) to make a 10 mM CFDA-SE stock (store at −20◦ C). Add 10 µl of this to 990 µl of PBS to make a 100 µM stock just before use; use immediately.

DAB DAB (Sigma) is a substrate of HRP that gives a brown product. It comes in various forms from various vendors. Keep in mind that the powder is extremely toxic so try to avoid that form. DAB is sold in various forms, from pellets to dissolve in water to ready-to-use solutions. The final concentration is 1 mg/ml. NOTE: Alternative chromogens are available to obtain different colors from HRP or to be used with different enzymes. One example is 9 ethyl-carbazole, which produces a red precipitate (Jung et al., 2007).

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Fluorescent vascular dyes The bone marrow vasculature can be easily observed if the mouse is injected with a fluorescent dye that persists in the circulation long enough without extravasating. FITC or rhodamine dextran are among the most commonly used vascular dyes (Cavanagh et al., 2005). Angiosense probes from Visen Medical are expensive but good alternatives to generate signal especially in the near-infrared region of the light spectrum (Montet et al., 2007). The bone marrow cavity has been visualized using dyes that do extravasate and flood the bone marrow (Cavanagh et al., 2005).

Ketamine/xylazine cocktail To a bottle of 10 ml Ketamine HCl (Henry Schein) 50 mg/ml (500 mg) add 750 µl xylazine (Henry Schein) 100 mg/ml (75 mg). Shake well. Store the bottle up to 3 months at room temperature in the dark. NOTE: Ketamine is a recreational drug and has to be purchased under license. This drug must be kept under lock and key, and all usage must be documented in a log book.

Long-term culture medium H5100 for human or M5300 for murine cells (StemCell Technologies) Just before using add: Penicillin/streptomycin (Cellgro, no. 30-001-CI, diluted to 1 in 500 in the medium) Hydrocortisone (StemCell Technologies) to a final concentration of 10−6 M Use immediately after adding supplements.

Methylcellulose-containing medium Use methylcellulose-containing medium H4435 for human cells and M3434 for murine cells, both from StemCell Technologies. Store the original bottles and the aliquots of medium frozen. Thaw the bottle of medium at room temperature (not at 37◦ C, for better growth factors preservation).

Vortex vigorously Prepare 3-ml aliquots Store the aliquots at −20◦ C until ready to use them Paraformaldehyde Prepare a stock of paraformaldehyde (PFA) up to 12% (w/v) by dissolving the powder (Sigma, stored at 4◦ C) in PBS without Ca or Mg, add a few drops of NaOH to reach pH 7.5 and heat up to 70◦ C while stirring. At higher temperatures the PFA breaks into formaldehyde, which is not as stable. Aliquot and freeze the stock solution. Thaw each aliquot, dilute to 3% and use for 1 to 2 weeks if stored at 4◦ C.

COMMENTARY Background Information

Analysis of the HSC Niche

The concept that stem cells, and in particular HSC, are regulated not only by cellautonomous mechanisms but also by a complex network of signals generated or conveyed by their specialized bone marrow microenvironment was proposed some decades ago, but only recent findings have allowed the identification of some HSC niche components and of

some of the molecular mechanisms regulating HSC-niche interactions. Strong evidence suggests that the osteoblasts in the bone marrow are a key HSC niche component. Involved in bone development, mineralization, and remodeling, osteoblasts also produce growth factors supporting HSC growth (Taichman et al., 2000). There is a direct correlation between the number of osteoblasts and the number

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of HSC (Calvi et al., 2003; Zhang et al., 2003). Molecules described to have a role in osteoblast-HSC cross-talk include: Jagged1 and Notch (Calvi et al., 2003), N-cadherin (Zhang et al., 2003), angiopoietin and Tie2 (Arai et al., 2004), and osteopontin (Nilsson et al., 2005; Stier et al., 2005). Homing of HSC to the bone marrow and engagement of the endosteal region have been shown and are known to be necessary in bone marrow transplant settings in order to lead to engraftment and bone marrow and peripheral blood reconstitution (Adams et al., 2006). Not only osteoblasts, but also osteoclasts have been proved to interact with HSC (Kollet et al., 2006). HSC have been observed in direct proximity of osteoblasts and also next to capillaries in marrow and spleen sections (Arai et al., 2004; Kiel et al., 2005). Leukemic cell lines and hematopoietic cells were injected into mice and observed to be rolling and homing in specific areas of marrow vasculature, where they remained up to 70 days later (Sipkins et al., 2005). Stromal cells located around bone marrow sinusoids or close to the endosteum have recently been indicated as the cells responsible for directing HSC homing by producing the chemokine stromal-derived factor 1 (SDF-1; Sugiyama et al., 2006). Moreover, the nervous system is known to reach the bone marrow and to influence the ability of HSC to engage the niche and be mobilized (Katayama et al., 2006) Even though these recent developments have started to shed light on the complex characteristics of the HSC niche, still little is known about the nature of its components and the molecular mechanisms of their interactions with HSC. Immunofluorescence and immunohistochemistry Theoretically it is possible to stain tissue sections for any antigen of interest to determine its localization and even quantify its abundance within the tissue or in different experimental conditions. Practically, the availability of highly specific antibodies is the limiting factor when planning immunostaining, and even though it is relatively straightforward to generate new polyclonal antibodies by immunizing rabbits with peptides from the antigen of interest, different antigens will have different immunizing activity, plus the peptides used for the immunization are not necessarily the most readily accessible in the tissue. It goes beyond the aims of this unit to present methods for the generation and testing of new antibodies or various treatments that can be performed in order to unmask antigens in tissue

sections. For more details on these topics see, for example, Lane and Harlow, 1999. Examples of well characterized antibodies that have been used so far in HSC niche studies are antiJagged1, osteopontin (Calvi et al., 2003) and N-cadherin (Zhang et al., 2003) to visualize osteoblasts, PECAM/CD31 to visualize vasculature (Sipkins et al., 2005), SLAM (CD150, CD48 and CD41, Kiel et al., 2005) and N cadherin (Zhang et al., 2003) to visualize HSC. The use of these antibodies allows evaluation for example of osteoblasts, HSC, and vessels number in the bone marrow of test and control mice. Intravital microscopy Intravital microscopy is becoming more popular as the best technique to generate a multidimensional view of cells interacting within a tissue (Iga et al., 2006; Kuebler et al., 2007; Soon et al., 2007). In the bone marrow, two-photon and confocal imaging have been used to observe memory T cells interacting with dendritic cells (Cavanagh et al., 2005; Mazo et al., 2005) and leukemic and hematopoietic cells extravasating and homing to perivascular space (Sipkins et al., 2005). This kind of analysis is excellent also to produce three-dimensional maps of expression of particular reporters (Runnels et al., 2006). Most intravital bone marrow imaging can be performed with confocal microscopes, but two-photon microscopy allows deeper imaging because it produces images with less noise compared to confocal microscopy (Zipfel et al., 2003). Homing and lodging The ability of HSC to reconstitute the bone marrow and peripheral blood of transplanted recipients relies on many factors. Before starting to self-renew and to give rise to a differentiating progeny, HSC have to find their way to the bone marrow (homing) and stably engage the niche (engraftment; Adams and Scadden, 2006). It is important to determine whether transplant failure is due to a stem cell-intrinsic defect, such as inability to self-renew, or to defects in the HSC-niche interaction. Homing and lodging assays allow such discrimination by assessing HSC performance soon after transplant. There is no universal agreement on the terminology to use when describing niche engagement by transplanted HSC and often homing and lodging assays are performed in different ways, but generally the homing assay is performed using lethally irradiated recipients, while the lodging

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assay is performed using nonirradiated recipients (compare for example Nilsson et al., 2001; Yang et al., 2007). In vitro culture of hematopoietic progenitor cells Historically CFU-C (colony-forming unit in culture), CAFC (cobblestone area forming cells), and LTC-IC (long-term culture initiating cell) assays were used as a read out of HSC number (Sutherland et al., 1990; Breems et al., 1994; Bouzianas, 2003; van Os et al., 2004). CFU-C assay used to compare HSC harvested from wild-type and microenvironment mutant mice can give an indication of the number and type of hematopoietic progenitors present in the mice, reflecting the ability of their bone marrow microenvironment to support hematopoiesis. There is a lot of debate on the value of these assays as a readout of HSC number, and the general consensus is that in vivo assays such as limiting dilution transplants are the best way to confirm in vitro data and give an indication of HSC number (van Os et al., 2004). The Stem Cell Technology Web site (http://www.stemcell.com/) contains detailed descriptions for the set up of all colony formation-based assays. The in vitro assays can be performed not only with murine but also with human hematopoietic cells and stroma, and are therefore an important component of studies on human HSC niche. Moreover, the possibility to transduce stroma cell lines with various constructs allows a first analysis of the molecular mechanisms regulating HSC-stroma interactions before proceeding to the generation of the appropriate transgenic mice.

Critical Parameters

Analysis of the HSC Niche

Several epifluorescence microscopes will be of sufficient quality to check the efficiency of staining and acquire images at low magnification. The use of a confocal microscope (e.g., Zeiss LSM series) is recommended to gain much greater detail by increasing resolution and diminishing background noise. A confocal image will always be more accurate than a simple epifluorescence picture, but because it allows a much higher control of the acquisition process, it is important to receive appropriate training in confocal microscopy before getting started. Some of the most common mistakes, such as using a too large pinhole (and thick optical slice), incorrect laser power, or inappropriate gain can determine the generation of misleading data, especially when analyzing

colocalization of markers or expression levels (see, for example, Pawley, 1995). When performing in vivo imaging experiments it is necessary to make sure the fluorophores used are sufficiently bright to be detected and that absorption and emission spectra are sufficiently far apart to be easily distinguished. When performing homing and lodging assays it is important to include a sufficient number of recipient mice in the test and control groups in order to generate statistically significant data. When the difference between the mean of two groups is known up front, it is possible to calculate the sample size (number of recipients) that will generate statistically significant data. In most cases the whole objective of the experiment is to find out such difference, therefore it is recommended to use between five and ten recipient mice per group. It is essential to work in perfectly sterile conditions when preparing long-term cultures of stroma/hematopoietic cells in order to avoid contamination.

Troubleshooting If it is not possible to visualize the central vein during in vivo imaging of the calvarium it is possible that the mouse head is tilted and it is advisable to re-adjust its position. If the mouse has already been imaged previously it is possible that scar tissue is generating enough light scatter to completely impair observation of the vein. Inconsistent results obtained when performing the homing assay with lineagedepleted cells is possibly due to variability intrinsic to the lineage depletion process. In this case it might be better to perform the assay using whole bone marrow monocytes and differentiate between lineage-positive and negative cells while analyzing the recipients’ bone marrow.

Anticipated Results When observing bone marrow vasculature in vivo the central vein of the calvarium appears as a wide, straight vessel, from which smaller vessels depart. Winding and relatively wide vessels depart from the central vein underneath the coronal suture. These vessels also are site of origin for bone marrow capillaries. When performing homing assay by injecting 5 million total bone marrow cells per recipient, typically 0.5% to 5% of the recipient cells observed will be labeled, and can be

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further subdivided between lineage positive or negative cells. When performing lodging assay injecting 500,000 Lin− wild-type cells into recipients, expect to observe about twelve cells per serially sectioned femur.

Time Considerations The definitive readout on HSC function, either cell-intrinsic or cell-extrinsic regulated, is obtained with bone marrow transplants, which require from 12 weeks to several months in order to be completed. The assays described in this unit are relatively shorter. The preparation of bones for histological analysis takes between a few hours to 5 days. A few hours are sufficient for an experienced person to cut sections from a number of blocks, and the typical immunofluorescence/immunohistochemistry staining will last from a few hours to a day (sometimes split by an overnight incubation). The preparation of cells for most of the other assays can be among the most time-consuming procedures, with sorting of HSC taking the better part of a day. In vivo imaging sessions should not last more than 3 to 4 hr each in order not to harm the mouse. The homing assay requires a waiting time of 4 to 24 hr between irradiation and injection and 6 hr after the injection, so it is recommended to irradiate the recipients in the evening and perform the whole assay the following day. The lodgment assay requires less time than the homing assay, but follows the schedule of bone embedding, sectioning and mounting, for the analysis of the results. CAFC/LTC-IC assay requires a long time to reach its end, but can easily be set up with 1 day of work to prepare the stroma and 1 day of work 3 to 5 weeks later to seed the HSC. About 1 hr should be sufficient to score a 96-well plate for CAFC or LTC-IC and evaluate the data with L-Calc.

Acknowledgements We thank the following people for their advice: Dr. Ernestina Schipani and Dilani Rosa on histology and immunohistochemistry methods, Professor Charles Lin and Juwell Wu on in vivo imaging methods, Dr. Maria Toribio on human HSC, Dr. Gregor Adams and Ian Alley on homing and lodgment assays, Dr. Louise Purton on colony formation assays, Mehron Puoris’haag and Simon Broad on suppliers of reagents. Dr. Aparna Venkatraman helped in revising the manuscript and Chris Shamburgh provided

administrative assistance. Dr. Lo Celso has been funded by the European Molecular Biology Organization and the Human Frontiers Science Program.

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Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18.

Kitajima, K., Tanaka, M., Zheng, J., Yen, H., Sato, A., Sugiyama, D., Umehara, H., Sakai, E., and Nakano, T. 2006. Redirecting differentiation of hematopoietic progenitors by a transcription factor, GATA-2. Blood 107:1857-1863. Kollet, O., Dar, A., Shivtiel, S., Kalinkovich, A., Lapid, K., Sztainberg, Y., Tesio, M., Samstein, R.M., Goichberg, P., Spiegel, A., Elson, A., and Lapidot, T. 2006. Osteoclasts degrade endosteal components and promote mobilization of hematopoietic progenitor cells. Nat. Med. 12:657-664. Kuebler, W.M., Parthasarathi, K., Lindert, J., and Bhattacharya, J. 2007. Real-time lung microscopy. J. Appl. Physiol. 102:1255-1264. Lane, D. and Harlow, E. 1999. Using Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Woodbury, NY.

Analysis of the HSC Niche

Mazo, I.B., Honczarenko, M., Leung, H., Cavanagh, L.L., Bonasio, R., Weninger, W., Engelke, K., Xia, L., McEver, R.P., Koni, P.A., Silberstein, L.E., and von Andrian, U.H. 2005. Bone marrow is a major reservoir and site of recruitment for central memory CD8+ T cells. Immunity 22:259-270.

Runnels, J.M., Zamiri, P., Spencer, J.A., Veilleux, I., Wei, X., Bogdanov, A., and Lin, C.P. 2006. Imaging molecular expression on vascular endothelial cells by in vivo immunofluorescence microscopy. Mol. Imaging 5:31-40. Scadden, D.T. 2006. The stem-cell niche as an entity of action. Nature 441:1075-1079. Schofield, R. 1978. The relationship between the spleen colony-forming cell and the haemopoietic stem cell. Blood Cells 4:7-25. Shizuru, J.A., Negrin, R.S., and Weissman, I.L. 2005. Hematopoietic stem and progenitor cells: Clinical and preclinical regeneration of the hematolymphoid system. Annu. Rev. Med. 56:509-538. Sipkins, D.A., Wei, X., Wu, J.W., Runnels, J.M., Cote, D., Means, T.K., Luster, A.D., Scadden, D.T., and Lin, C.P. 2005. In vivo imaging of specialized bone marrow endothelial microdomains for tumour engraftment. Nature 435:969-973.

Lo Celso, C. and Scadden, D.T. 2007. Isolation of Hematopoietic Stem Cells from Bone Marrow. In Journal of Visualized Experiments Issue 2.

Soon, L., Braet, F., and Condeelis, J. 2007. Moving in the right direction-nanoimaging in cancer cell motility and metastasis. Microsc. Res. Tech. 70:252-257.

Lo Celso, C., Prowse, D.M., and Watt, F.M. 2004. Transient activation of beta-catenin signalling in adult mouse epidermis is sufficient to induce new hair follicles but continuous activation is required to maintain hair follicle tumours. Development 131:1787-1799.

Stier, S., Ko, Y., Forkert, R., Lutz, C., Neuhaus, T., Grunewald, E., Cheng, T., Dombkowski, D., Calvi, L.M., Rittling, S.R., and Scadden, D.T. 2005. Osteopontin is a hematopoietic stem cell niche component that negatively regulates stem cell pool size. J. Exp. Med. 201:1781-1791.

2A.5.30 Supplement 3

Current Protocols in Stem Cell Biology

Sugiyama, T., Kohara, H., Noda, M., and Nagasawa, T. 2006. Maintenance of the hematopoietic stem cell pool by CXCL12-CXCR4 chemokine signaling in bone marrow stromal cell niches. Immunity 25:977-988. Sutherland, H.J., Lansdorp, P.M., Henkelman, D.H., Eaves, A.C., and Eaves, C.J. 1990. Functional characterization of individual human hematopoietic stem cells cultured at limiting dilution on supportive marrow stromal layers. Proc. Natl. Acad. Sci. U.S.A. 87:3584-3588. Sutherland, H.J., Eaves, C.J., Lansdorp, P.M., Thacker, J.D., and Hogge, D.E. 1991. Differential regulation of primitive human hematopoietic cells in long-term cultures maintained on genetically engineered murine stromal cells. Blood 78:666-672. Taghon, T.N., David, E.S., Zuniga-Pflucker, J.C., and Rothenberg, E.V. 2005. Delayed, asynchronous, and reversible T-lineage specification induced by Notch/Delta signaling. Genes Dev. 19:965-978. Taichman, R.S., Reilly, M.J., and Emerson, S.G. 2000. The Hematopoietic microenvironment: Osteoblasts and the hematopoietic microenvironment. Hematology 4:421-426. van Os, R., Kamminga, L.M., and de Haan, G. 2004. Stem cell assays: Something old something

new, something borrowed. Stem Cells 22:11811190. Wolf, N.S. 1974. Dissecting the hematopoietic microenvironment. I. Stem cell lodgment and commitment, and the proliferation and differentiation of erythropoietic descendants in the S1-S1d mouse. Cell Tissue Kinet. 7:89-98. Yang, L., Wang, L., Geiger, H., Cancelas, J.A., Mo, J., and Zheng, Y. 2007. Rho GTPase Cdc42 coordinates hematopoietic stem cell quiescence and niche interaction in the bone marrow. Proc. Natl. Acad. Sci. U.S.A. 104:5091-5096. Zhang, J., Niu, C., Ye, L., Huang, H., He, X., Tong, W.G., Ross, J., Haug, J., Johnson, T., Feng, J.Q., Harris, S., Wiedemann, L.M., Mishina, Y., and Li, L. 2003. Identification of the haematopoietic stem cell niche and control of the niche size. Nature 425:836-841. Zhu, J., Garrett, R., Jung, Y., Zhang, Y., Kim, N., Wang, J., Joe, G.J., Hexner, E., Choi, Y., Taichman, R.S., and Emerson, S.G. 2007. Osteoblasts support B lymphocyte commitment and differentiation from hematopoietic stem cells. Blood 109:3706-3712. Zipfel, W.R., Williams, R.M., and Webb, W.W. 2003. Nonlinear magic: Multiphoton microscopy in the biosciences. Nat. Biotechnol. 21:1369-1377.

Somatic Stem Cells

2A.5.31 Current Protocols in Stem Cell Biology

Supplement 3

Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

UNIT 2A.6

Alexander Medvinsky,1 Samir Taoudi,1 Sandra Mendes,2 and Elaine Dzierzak2 1 2

Institute for Stem Cell Research, University of Edinburgh, Edinburgh, United Kingdom Erasmus Medical Center, Department of Cell Biology, The Netherlands

ABSTRACT Hematopoietic development begins in several locations in the mammalian embryo: yolk sac, aorta-gonad-mesonephros region (AGM), and the chorio-allantoic placenta. Generation of the most potent cells, adult definitive hematopoietic stem cells (HSCs), occurs within the body of the mouse embryo at midgestation in the AGM region. Similarly, at the equivalent developmental time in the human embryo, the AGM region has been shown to contain multipotent progenitors. Hence, the mouse embryo serves as an excellent model to study hematopoietic development. To further studies on the ontogeny of the adult hematopoietic system, the focus of this unit is on the experimental methods used in analysis of the AGM region. Curr. Protoc. Stem Cell Biol. 4:2A.6.1-2A.6.25. C 2008 by John Wiley & Sons, Inc.
Keywords: developmental hematopoiesis r hematopoietic stem cells r embryo r AGM r lineage differentiation

INTRODUCTION Development of the hematopoietic system is a complex process occurring in several embryonic locations. Since there is a high degree of conservation between the hematopoietic systems of mouse and humans, the mouse is an excellent experimental model for the study of blood development. Here the focus is on experimental approaches in the mouse embryo facilitating the analysis of the aorta-gonad-mesonephros (AGM) region, a tissue central to the development of the adult hematopoietic system. Protocols describe how to dissect the AGM region (see Basic Protocol 1 and Support Protocols 1 and 2), prepare a cell suspension (see Basic Protocol 2), culture the cells (see Basic Protocol 3), isolate various cell populations (see Basic Protocol 4), and analyze AGM cell lineage potential in various hematopoietic (see Basic Protocols 5 and 6, Support Protocols 3 and 4), endothelial (see Basic Protocol 7 and Support Protocol 5), and mesenchymal differentiation assays (see Basic Protocol 8 and Support Protocols 6, 7, 8, and 9). These methods will be useful for those who study molecular and cellular mechanisms of hematopoietic development, with particular focus on the development of adult-type (definitive) hematopoietic stem cells (HSCs), and also for those who are interested in the analysis of the relationship between hematopoietic and non-hematopoietic lineages.

DISSECTION OF MOUSE EMBRYONIC TISSUES FROM DAY 9 TO 12 MOUSE EMBRYOS

BASIC PROTOCOL 1

Several embryonic sites are involved in hematopoiesis: (1) intra-body sites: the para-aortic splanchnopleura (Pa-Sp), which by embryonic day 9 develops into the aorta-gonadmesonephros (AGM) region and the liver; (2) extra-body sites: the yolk sac and placenta; and (3) blood vessels: the umbilical and vitelline vessels that connect the embryo body to the placenta and yolk sac, respectively. Somatic Stem Cells Current Protocols in Stem Cell Biology 2A.6.1-2A.6.25 Published online January 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a06s4 C 2008 John Wiley & Sons, Inc. Copyright


2A.6.1 Supplement 4

Sterile preparation of cells for in vitro and in vivo assays is recommended. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

Materials Pregnant female mice of chosen background strain 70% ethanol Medium I (medium for embryo collection; see recipe) Medium II (medium for dissections; see recipe) Surgical scissors (two pairs treated with 70% alcohol) Fine, straight watchmaker’s forceps (two pairs) 60 × 15–mm and 35 × 15–mm plastic tissue culture petri dishes 150-W cold light source equipped with double gooseneck fiber-optic system Dissection microscope (magnification range from 7× to 40× with a flat, black background stage; Leica, Zeiss, or Olympus) Fine, curved watchmaker’s forceps Dissection needles: sharpened tungsten wire 0.375-mm diameter (Agar Scientific Ltd.) attached to metal holders typically used for bacterial culture inoculation (alternatively, 29-G needles attached to micro-fine insulin syringes, e.g., U-100, Beckton-Dickinson) Device for sharpening dissection needles (an electrolytic device for sharpening tungsten needles described in Hogan and Beddington, 2002, or alternatively, a sharpening stone) NOTE: For dissections, always use room temperature solutions. When necessary to maintain sterility during tissue isolation, wash dissection tools with 70% alcohol and wipe with a tissue. Excess blood should first be removed from tools by wiping with distilled water.

Collect embryos 1. Sacrifice pregnant females by cervical dislocation at the desired day/stage of gestation. The AGM region can be dissected from embryos between E10.5 and E13.5 of development.

2. Wash the abdomen of the animal with 70% ethanol. Make a transverse incision with scissors and open the mesenteric layer underlying the skin by pulling the skin apart with fingertips. The mesenteric layer should be kept intact at this stage.

3. Using another pair of scissors make a transverse incision through the mesenteric layer at the level of the abdomen. Avoid internal cuts so as not to injure internal tissues, especially the digestive tract. 4. Locate the uterus and, using straight forceps, pull one horn of the uterus out of the abdomen. Separate it from the mesenteric tissue with the scissors; continue along the second horn of the uterus. Cut close to the uterus to maximally remove mesenteric fat adjacent to it. Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

5. Place the uterus in a 60 × 15–mm tissue culture dish containing medium I. Continue removing the remaining adipose tissue and then move the uterus into a clean culture dish containing medium I. 6. Position the gooseneck 150-W light source to provide clear illumination of the contents of the culture dish. Visualize the uterus at low magnification (7× to

2A.6.2 Supplement 4

Current Protocols in Stem Cell Biology

Figure 2A.6.1 Dissection of an E11 mouse conceptus. (A) Embryo with chorionic membrane being removed. (B) Separation of the placenta (PL) from the yolk sac which envelops the embryo. (C) After the yolk sac (YS) is disrupted, it no longer envelops the embryos but is still attached to it through the vitelline vessels. (D) The umbilical cord (UC) is seen connected to the embryo body at one end and the disrupted yolk sac is visible at the head of the embryo. (E) The head and upper region of the embryo body to the forelimbs is severed from the trunk of the embryo. (F) The dorsal tissues, neural tube (NT), and somite tissue (ST) are dissected away from the embryo trunk region. (G) After removal of the dorsal tissues, the dorsal aorta (Ao) is visible along the midline on a view of the dorsal trunk. (H) On the ventral trunk, the umbilical vessels (UC) are visible. The liver (FL) is seen as the pink tissue just above the umbilical cord. (I) A dorsal trunk region view showing the body walls (BW) lateral to the AGM have been dissected away. In this dorsal view, the urogenital ridges (UGR) are laterally juxtaposed to the dorsal aorta (Ao). (J) Ventral view of the AGM (only a small part of the UGR is visible) with overlying ventral tissues; stomach (ST) and liver (FL) and the fetal liver (FL) is still attached. (K) Crudely dissected and separated fetal liver (left) and AGM. (L) Cleanly dissected AGM region viewed from the ventral aspect. Ao = aorta (DA). Urogenital ridges located lateral to the Ao are clearly visible, with the genital ridge/developing gonads overlaying the pronephros and mesonephros (embryonic kidney).

8×) under a dissection microscope. Using two pairs of fine straight forceps, open the muscular wall layer of the uterus and isolate deciduas with embryos (Fig. 2A.6.1A). 7. Then with small grasps of the forceps, remove the decidua and Reichert’s membrane, which is the thin tissue layer surrounding the yolk sac (Tavian and Peault, 2005). It is best not to rupture the yolk sac membrane. The maintenance of yolk sac integrity, as well as placenta localization, allows a ready recognition of the vitelline and umbilical vessels connecting the extraembryonic tissues to the embryo. If experimentation with the embryonic cells requires long-term in vitro culture, the dissections should be performed under a microscope placed in a horizontal flow cabinet.

8. During these manipulations, gently transfer the embryos by placing curved forceps under the embryo to support and move it into clean culture dishes containing medium I to wash away maternal blood contamination. Close the forceps only very loosely around the embryo, so as not to damage or break the extraembryonic membranes.

Isolate placenta 9. Dissect placentas free of the embryo by gently separating it from the yolk sac and tightly closing the straight fine forceps around the umbilical vessels at the junction

Somatic Stem Cells

2A.6.3 Current Protocols in Stem Cell Biology

Supplement 4

with the placenta to cut connection (Fig. 2A.6.1B,D). Remove any parts of the yolk sac that remain attached from the placenta. Also, remove the maternal decidua.

Isolate yolk sac 10. Grasp the yolk sac with the fine-tipped forceps and tear open this tissue to reveal the embryo. Remove the embryo from the yolk sac by closing the forceps tightly around the vitelline vessels and severing them at their connection with the yolk sac (Fig. 2A.6.1C). The amniotic sac should now be the only membrane remaining around the embryo, although this thin and almost transparent membrane may have broken open during the previous dissections.

Isolate vitelline and umbilical vessels 11. Obtain the vitelline and umbilical arteries by severing their connection to the embryo body proper with fine scissors or the tight closure of the fine forceps at this junction (Fig. 2A.6.1D,H). Isolate fetal liver and AGM 12. Lay the whole embryo onto one side. Dissect the head region away by placing one dissection needle dorsally and one dissection needle ventrally to direct the cutting action just above the forelimbs (Fig. 2A.6.1E). Use dissection needles for the isolation of the AGM and liver. Adjust the microscope to a slightly higher magnification. Hold one needle in each hand and gently place one needle in the area where cutting is desired to immobilize the embryo. Place the other needle on the other side of the region to be cut and slowly move it along the holding needle so that the crossing of the needles results in a cutting action. For the most precise dissections, only small areas are cut with each action.

13. Similarly, use the needles to cut across and remove the tail region, just below the hind limbs. 14. Continue with the isolation of the AGM from the trunk region by removing dorsal tissues, including the somites and neural tube. Place one needle within the somite region to immobilize the trunk tissue. With repetitive crossing and cutting actions moving from one end of the trunk region to the other, remove the dorsal tissue (Fig. 2A.6.1F). Careful small cutting actions are recommended to maintain the integrity of the dorsal aorta. The blood within the dorsal aorta serves as a landmark for the AGM (Fig. 2A.6.1G).

15. Since not all somite tissue will be removed, turn the trunk of the embryo slightly so that the ventral side is facing upwards (Fig. 2A.6.1H). Place the needles below the gonad-mesonephros and with small crossing and cutting actions proceed along the anterior-posterior axis to remove the rest of the somite tissue (Fig. 2A.6.1I). 16. Dissect the remaining trunk region (Fig. 2A.6.1J) more finely to remove the ventral tissues: liver and gastrointestinal (GI) tract (Fig. 2A.6.1K). To do this, place one needle in the connective tissue between the AGM and the heart. Place the other needle a short distance posteriorly in this connective tissue. Cross the needles and cut. Continue to move posteriorly, crossing and cutting with the needles until the ventral tissues are removed. Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

The dorsal aorta and laterally located gonad-mesonephros can now be seen (Fig. 2A.6.1L). A finer dissection of the liver can be performed to remove connective and GI tissue.

2A.6.4 Supplement 4

Current Protocols in Stem Cell Biology

GENERATING MOUSE EMBRYOS To obtain embryos, introduce two adult female mice (8 to 16 weeks old) into a cage containing one adult male mouse (8 to 50 weeks old) in the late afternoon. Early in the morning of the next day, check female mice for the presence of a vaginal plug. If a plug is found, move the female to another cage and note the date of plug discovery on the cage card. This is considered to be embryonic day 0.5 (E0) or 0.5 days post-coitum (dpc).

SUPPORT PROTOCOL 1

Isolated embryonic tissues are used for various hematopoietic studies. When used for in vivo transplantations, it is necessary that the donor embryonic cells contain a marker unique from the recipient. Often, a transgene (LacZ, GFP) is used as the genetic marker of the donor embryonic cells (deBruijn et al., 2000; North et al., 2002), although other markers are available, such as the Y chromosome marker (if embryos are typed for sex and male cells are injected into female recipients; Muller et al., 1994) or the Ly5.1/5.2 alleles (Bertrand et al., 2005). Since maternal blood cells are a source of contamination during the dissection of embryos, using a paternally derived transgene or the Y chromosome as the donor embryonic cell marker in transplantations is advantageous, in that it ensures that engraftment is from embryo-derived and not maternally derived cells.

STAGING EMBRYOS Embryos within a litter are staged by counting somite pairs (sp), examining eye pigmentation, and noting the shape of the limb buds. Since the embryos within a single litter can vary by as much as 0.5 days in gestation, precise somite counts assure that embryonic tissues to be used for an experiment will be developmentally similar.

SUPPORT PROTOCOL 2

For better contrast, a dissection microscope with a magnification range of 7× to 40×, a black background stage, and a 150-W cold light source equipped with a double gooseneck fiber-optic system is used to illuminate the embryos from the side (at 10× to 15× magnification). E8 to E8.5 embryos have 1 to 7 sp; E8.5 to E9 embryos have 8 to 14 sp; E9 to E9.5 embryos have 13 to 20 sp, and E9.5 to E10 embryos have 21 to 30 sp. Embryos of 30 to 35 sp are considered early E10, 36 to 37 sp mid-E10, and 38 to 40 sp late E10. At E11, somite pairs are >40, the eye pigmentation ring is closing, and the limb buds are rounded with the beginning of internal digital segmentation.

PREPARATION OF CELL SUSPENSION FROM TISSUES OF MIDGESTATION MOUSE EMBRYOS

BASIC PROTOCOL 2

Prior to transplantation, flow cytometric analysis/sorting, or the in vitro culture of embryonic tissues, it is necessary to produce a single-cell suspension. This is accomplished in two phases: step one involves enzymatic digestion of the dissected organ; and step two involves the mechanical disruption of the organ by gentle pipetting. Once a cellular suspension has been produced from the desired organ, various assays can be used to investigate and manipulate its functional properties.

Materials Collagenase type I (see recipe) Embryonic tissues (Basic Protocol 1) Medium II (see recipe) Medium III (see recipe), room temperature and ice cold 10-ml round-bottom transparent polystyrene tubes (Sterilin) 37◦ C water bath with shaking Vacuum aspirator NOTE: Keep cell suspensions strictly on ice.

Somatic Stem Cells

2A.6.5 Current Protocols in Stem Cell Biology

Supplement 4

1. Thaw and dilute collagenase type I stock 1:20 in medium II. 2. Add 0.5 to 1.5 ml (dependent upon the mass of tissue) diluted collagenase type I in a 10-ml round-bottom tube. A volume of 1 ml of 0.12% collagenase will disperse about ten embryonic E11.5 AGM regions when incubated 1 hr at 37◦ C.

3. Place tissues in 10-ml tubes containing diluted collagenase type I and incubate 30 to 90 min, depending on tissue type and mass, in a 37◦ C water bath with slow shaking. Large tissues such as placenta and E12.5 liver should be cut in several pieces (incubate placenta for 90 min).

4. Following incubation, wash the tissues by adding room temperature medium III to bring volume up to 5 ml and centrifuging 5 min at 300 × g, room temperature. 5. Carefully remove supernatant and flick all tubes to disperse the cells, add 1 ml of ice-cold medium III to each tube and place on ice. 6. Without delay, gently pipet tissues up and down (triturate) ∼25 times using a pipettor with a large-bore tip. For large tissues, first use a tip with the end cut. Avoid bubbles as they decrease cell viability. Do not try to make a true single cell suspension, as this will increase the number of dead cells.

7. Sediment large cell aggregates by positioning the tube vertically on ice for 1 to 3 min. Transfer the cell suspension into a new 10-ml tube and keep it on ice. Do not use 15-ml conical tubes as cell suspension cannot be collected from them using a 1-ml syringe.

8. Centrifuge cells at 300 × g, 4◦ C. To ensure that all cells are sedimented, use the following formula to determine the length of centrifugation: no. of min of centrifugation = no. of ml in 10-ml tube + 2 min. 9. Carefully remove the supernatant using a vacuum aspirator with the tip of the pipet touching only the surface of the solution to avoid disturbing the pellet. Stop aspirating when the 0.5-ml mark on the tube is reached. Gently resuspend the cells and leave them on ice. The cells have been kept on ice for up to 6 hr with preservation of hematopoietic function, however, once placed on ice, the cells should be used as soon as possible. BASIC PROTOCOL 3

Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

EXPLANT CULTURE OF EMBRYONIC TISSUES The in vitro culture of whole tissues allows for the autonomous growth of the tissue in the absence of cellular exchange with other tissues that is occurring through circulation or interstitial migration. A closed circulatory system is established in the mouse conceptus at E8.5 (9 sp stage). Explant cultures are particularly useful for testing the effects of exogenously added growth factors. These cultures demonstrate that any stem cells found at a later time point are derived from the explanted tissue, and not from cells that have migrated in.

Materials Explant medium: myeloid long-term culture medium (Stem Cell Technologies cat. no. M5300) supplemented with hydrocortisone succinate (Sigma) at a final concentration of 10−5 M Embryonic tissues (Basic Protocol 1) PBS or sterile water (Sigma)

2A.6.6 Supplement 4

Current Protocols in Stem Cell Biology

70% ethanol Collagenase type I (see recipe) Stainless-steel wire mesh supports (see recipe) 6-well tissue culture plates Straight and curved fine-tipped watchmaker’s forceps 0.65-µm membrane filters (Millipore Durapore) Scalpel blade NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. 1. Place a stainless-steel wire mesh support into the well of a 6-well culture plate. Fill the well with 5 ml of explant medium. Using the straight-tipped forceps, gently place a 0.65-µm membrane filter on top of the wire mesh, allowing it to absorb medium from one edge until it is completely wet. Adjust the medium level so that the filter is at the air-medium interface. 2. Using the curved-tipped forceps, place the embryonic tissues on top of the filter. Each filter can accommodate up to six individual tissues (e.g., E11.5 AGM regions) or fragments of large tissues (e.g., placenta).

3. To ensure appropriate humidity during culture, fill the empty wells of the culture plate with PBS or sterile water. Culture explants 2 to 3 days in a 37◦ C, 5% CO2 incubator. 4. Wearing gloves washed with 70% ethanol, pick up the filter with the forceps. Hold a scalpel with the other hand and gently scrape each tissue individually from the filter into a 10-ml tube. Place the tissue in 500 µl collagenase to make a single-cell suspension (see Basic Protocol 2) and place on ice.

PREPARATION OF EMBRYONIC CELLS FOR FLOW CYTOMETRY The following protocol is used for the processing of cellular suspensions of embryonic yolk sac, liver, placenta, and peripheral blood cells for flow cytometric analysis and sorting.

BASIC PROTOCOL 4

Materials Single-cell suspension from embryonic tissues (see Basic Protocol 2) FACS wash buffer: ice-cold 7% FBS/CMF-PBS (Sigma cat. no. D8537) Fc-block (anti-CD16/32 antibodies/Fc-γ III/II receptor) (BD Bioscience; Clone 2.4G2) Appropriate experimental antibodies (Table 2A.6.1) 7-Amino-actinomycin D (7-AAD; eBioscience; Table 2A.6.2) 40-µm nylon cell strainer (BD Falcon) 5.0-ml polystyrene tubes (BD Falcon) Refrigerated swing-out centrifuge Flow cytometer (e.g., FACSCalibur, BD Biosciences) Prepare cells 1. Prepare cellular suspensions from embryonic organs. 2. Remove large clumps by passing the cell suspension through a 40-µm nylon cell strainer.

Somatic Stem Cells

2A.6.7 Current Protocols in Stem Cell Biology

Supplement 4

Table 2A.6.1 List of Useful Primary Antibodies

Antigen

Clone

Isotype

Working concentration

Supplier

α4-Integrin

9C10

Rat IgG2α,κ

2.0 µg/ml

Pharmingen

AA4.1

AA4.1

Rat IgG2β,κ

2.0 µg/ml

eBioscience

c-Kit

2B8

Rat IgG2β,κ

2.0 µg/ml

Pharmingen

CD16/32

2.4G2

Rat IgG2β,κ

2.0 µg/ml

Pharmingen

CD34

RAM34

Rat IgG2α,κ

2.0 µg/ml

Pharmingen

CD41

MWReg30

Rat IgG1,κ

2.0 µg/ml

Pharmingen

CD45

30-F11

Rat IgG2β,κ

2.0 µg/ml

Pharmingen

Flk-1

Avas-12α

Rat IgG2α,κ

2.0 µg/ml

Pharmingen

Ly-5.1

A20

Mouse IgG2α,κ

2.0 µg/ml

eBioscience

Ly-5.2

104

Mouse IgG2α,κ

2.0 µg/ml

eBioscience

Mac-1

M1/70

Rat IgG2β,κ

2.0 µg/ml

Pharmingen

PECAM-1

MEC 13.3

Rat IgG2α,κ

2.0 µg/ml

Pharmingen

Sca-1

D7

Rat IgG2α,κ

2.0 µg/ml

eBioscience

Ter119

TER-119

Rat IgG2β,κ

2.0 µg/ml

eBioscience

Tie-2

TEK4

Rat IgG1,κ

2.0 µg/ml

eBioscience

VE-cadherin

11D4.1

Rat IgG2α,κ

6.0 µg/ml

Pharmingen

Table 2A.6.2 List of Useful Secondary Reagents

Reagent

Clonea

Working concentration

Supplier

7-AAD

n/a

0.5 µg/ml

eBioscience

Anti-rat IgG

Polyclonal

2.0 mg/ml

Southern Biotech

Mouse IgG2α,κ

G155-178

As appropriate

Pharmingen

Rat IgG1,κ

eBRG1

As appropriate

eBioscience

Rat IgG2α,κ

R35-95

As appropriate

Pharmingen

Rat IgG2β,κ

A95-1

As appropriate

Pharmingen

Streptavidin (fluorochromeconjugated)

n/a

0.2 µg/ml

Pharmingen

a n/a, not applicable.

3. Place 1 × 105 –106 cells in 100 µl of FACS wash buffer in a 5.0-ml polystyrene tube. 4. Add 100 µl of anti-CD16/32 antibody (5 µg/ml) to cells. Incubate 15 min on ice in the dark. This antibody binds high-affinity IgG Fc receptors and thus reduces background staining. Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

Stain cells 5. Add experimental antibody (diluted to the appropriate empirically determined concentration in FACS wash). Incubate cells 20 to 45 min on ice in the dark. 6. Add 1.0 ml FACS wash buffer and centrifuge tubes 5 min at 300 × g, 4◦ C (in the dark).

2A.6.8 Supplement 4

Current Protocols in Stem Cell Biology

Figure 2A.6.2 Sort criteria used to purify populations from the E11.5 AGM region. In the left panel, viable cells are identified on the basis of cell size (forward light scatter) and the exclusion of the nuclear stain 7-amino-actinomycin D (7-AAD). Viable cells in the gated region are then separated into endothelial (VE-cadherin+ CD45− ), hematopoietic (VEcadherin− CD45+ ), stem/progenitor (VE-cadherin+ CD45+ ), and non-endothelial/haematopoietic lineages (VE-cadherin− CD45− ) according to the differential plasma membrane expression of VE-cadherin and CD45 (right panel). According to this strategy, LTR-HSCs can be purified to a high frequency in the VE-cadherin+ CD45+ population (North et al., 2002; Taoudi et al., 2005).

7. Carefully aspirate supernatant with a manual pipet to minimize the risk of disturbing the pellet. 8. Proceed with secondary staining if required or continue to step 9. Generally, if fluorochrome-conjugated reagent is required to detect a primary antibody, the secondary reagent is added to the cell pellet that was resuspended in 100 µl FACS solution. Incubate 20 min on ice in the dark then wash cells as in step 6.

Final preparation 9. Resuspend cells in an appropriate volume of 0.5 µg/ml 7-AAD diluted in FACS wash. For sorting, resuspend cells at a concentration of 1 × 107 cells/ml and for analysis, 3–5 × 106 cells/ml. This can be determined empirically by testing the efficiency of dead cell detection in a mixture containing a known ratio of live to dead cells.

10. Analyze cells at a rate no >5000 events/sec. During sorting, acquire cells at a rate of ∼10,000 events/sec in FACS wash buffer. A typical example of the criteria used for both analysis and sorting of E11.5 AGM region cells expressing VE-cadherin and CD45 can be seen in Figure 2A.6.2. See Taoudi et al. (2005) for a discussion of marker expression by cells in E11.5 to E13.5 hematopoietic organs. The placement of gates for both cell analysis and cell sorting should be based on appropriate isotype control staining.

HEMATOPOIETIC (MYELOID) CLONOGENIC ASSAY This assay enables assessment of the number of progenitors or colony-forming cells (CFC, also known as CFU and CFU-C) in a single-cell suspension of embryonic tissues. The semi-solid/viscous nature of methylcellulose-based medium allows the differentiated cells produced by one progenitor cell to stay together as a distinct colony. This assay is not suitable to distinguish HSCs from hematopoietic progenitor cells.

BASIC PROTOCOL 5

Somatic Stem Cells

2A.6.9 Current Protocols in Stem Cell Biology

Supplement 4

Materials MethoCult GF M3434: complete methylcellulose-based medium with cytokines (SCF, IL-3, IL-6, erythropoietin) for detection of BFU-E, CFU-GM, CFU-G, CFU-M, and CFU-GEMM-formed colonies (100 ml, Stem Cell Technologies cat. no. 03434) Cell suspension 7% FBS/CMF-PBS (Sigma) DPBS, sterile 7-ml Bijou tubes (Sterilin) 2 and 10-ml syringes 18-G needles Neubauer hemacytometer 30- and 140-mm Petri dishes (non-adherent surface) 37◦ C, 5% CO2 incubator Inverted microscope Gridded 60-mm Petri dish Additional reagents and equipment for trypan blue staining (UNIT 1C.3) 1. Thaw MethoCult GF M3434 medium overnight at 4◦ C, mix well, and let stand for at least 5 min to allow bubbles to dissipate. 2. Dispense 2.3-ml aliquots of MethoCult GF M3434 into 7-ml Bijou tubes using a 10-ml syringe with 18-G needle (store at −20◦ C). 3. Count viable cells in the cell suspension using a Neubauer hemacytometer and trypan blue staining (UNIT 1C.3). 4. Adjust cell concentration for methylcellulose cultures in 7% FBS/CMF-PBS. Add 230 µl of cells to a 2.3-ml methylcellulose aliquot and vigorously mix. Let bubbles dissipate for several minutes. Typically, each plate is inoculated with 0.5 embryo equivalents of cells from the E11.5 AGM region (∼75,000 viable cells).

5. Transfer 1.1 ml of cells in methylcellulose medium into a 30-mm Petri dish with a nonadherent surface. Prepare two such dishes for each sample. 6. Place a maximum of eight 30-mm dishes into a 140-mm Petri dish and add one additional uncovered dish filled with sterile DPBS to prevent dehydration of cultures. Incubate 7 to 10 days in a 37◦ C, 5% CO2 incubator. 7. Score colonies under an inverted microscope. Place the individual culture dishes on the gridded 60-mm Petri dish to allow for a systematic scoring of colonies. Calculate average number of colonies. The expected distribution of CFU-C types from the E11.5 AGM region can be seen in Figure 2A.6.3A; the expected morphology and cellular composition of colonies can be seen in Figure 2A.6.3B. The optimal number of CFU-C colonies per 35-mm dish is between 30 and 60. This number normally provides sufficient data for statistical analysis and enables distinction between neighboring hematopoietic colonies. Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

There are a few differences in the colonies formed between embryonic and adult hematopoietic CFU-Cs. In the example of the AGM region, all colony types can be readily identified between 7 and 9 days of culture. In addition, CFU-mast derived from AGM region can present a CFU-GM-like morphology; therefore, time should be taken to practice the accurate classification of colony types.

2A.6.10 Supplement 4

Current Protocols in Stem Cell Biology

Figure 2A.6.3 Distribution and classification criteria of clonogenic colony forming units-culture (CFU-C) within the AGM region. (A) Distribution of CFU potential within one embryo equivalent (∼150,000 viable cells) of cells from the E11.5 AGM region. CFU-C identity is retrospectively ascribed following analysis of lineage potential using MethoCult medium (Stem Cell Technologies). (B) Criteria used for the classification of CFU-C identity: BFU-E produce erythroid cells in the presence of either macrophages or megakaryocytes; CFU-Mac, monocytes/macrophages; CFU-Mast, mast cells; CFUGM, granulocytes and monocytes/macrophages; CFU-GEMM, granulocytes, monocytes/macrophages, erythroid cells and megakaryocytes (Meg). Colony images (top panels), original magnification 40×; cytospin preparations (bottom panels), original magnification 630×.

LONG-TERM REPOPULATION ASSAY This assay enables the detection of definitive hematopoietic stem cells (HSCs). One HSC is capable of repopulating the entire hematopoietic system of an irradiated recipient, a property not processed by downstream CFU-Cs. Therefore, if the hematopoietic system of the recipient contains donor-derived cells after 3.5 months post-transplantation, or longer, then the injected cell suspension contained at least one HSC. Donor embryonic cells should be distinguishable from recipient blood cells. Here the Ly5.1/Ly5.2 system using mice on the C57Bl6 background is described. Ly5.2 is a wild-type pan-leukocytic CD45 allele, whereas Ly5.1 is a mutant CD45 allele. Commercially available antibodies can be purchased to distinguish Ly5.1- and Ly5.2-expressing cells.

BASIC PROTOCOL 6

Materials Adult recipient Ly5.1 C57Bl6 mice Ly5.2 C57Bl6 embryos CMF-PBS (Ca2+ /Mg2+ -free; Sigma) Adult Ly5.1/2 C57Bl6 mice bone marrow cells Mouse food Acidified water containing neomycin (Support Protocol 3)

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Supplement 4

200 mg/liter EDTA/PBS (Sigma) PharmLyse (BD Biosciences) or preferred red blood cell lysis buffer FACS wash buffer: 3% (v/v) FBS/CMF-PBS Anti-Ly5.1 and Ly5.2 monoclonal antibodies conjugated with alternative fluorochromes 137

Cesium irradiator (e.g., Gammacell GC40, MDS Nordion) Mice cages with heating pads Mouse holder with opening allowing extension of the tail 1-ml plastic syringes with 27-G needles 1.5-ml microcentrifuge tubes Swing-out centrifuge Flow cytometer (e.g., FACSCalibur, BD Biosciences)

1. Irradiate mice at 9.5 Gray with a 137 cesium irradiator. Preferably split irradiation into two doses with 3-hr interval as this allows mice to tolerate the irradiation better.

2. Prepare donor embryo cell suspensions (see Basic Protocols 1 and 2) in CMF-PBS and keep them on ice before transplantation. Harvest bone marrow carrier cells by flushing the femurs of adult Ly5.2/1 mice with 1.0 ml ice-cold FACS wash buffer (use a 1.0-ml syringe and 27-G needle). Count cells (2 × 104 nucleated cells should be prepared per recipient). Carrier bone marrow cells provide short-term rescue for irradiated mice before embryonic donor HSCs produce sufficient number of hematopoietic cells to rescue the animal in the long-term. Samples for transplantation should not contain more than 2% FBS as it may cause an undesirable immunological response. It is recommended to perform transplantations not later than within 3 hr after irradiation.

3. Place recipient mice in a cage on a heating pad and wait until their tail veins expand. 4. Place a warmed-up recipient mouse in a plastic mouse holder for intravenous injection of cells into the tail vein. 5. Mix the sample to be injected, e.g., by gently flicking the sample tube (avoid making bubbles). Fill a 1-ml syringe body with a 27-G needle with the cell suspension and slowly inject the cells into a lateral tail vein. Ensure that air bubbles are not injected into the vein. A transplanted volume of cells per mouse should not normally exceed 0.25 ml. The amount of cell suspension injected is dependent on the intended experiment. For example, a minimum of 1 equivalent of E11.5 AGM cells in 0.25 ml per recipient would be required to ensure successful reconstitution, while a 0.1 equivalent per recipient from explanted AGM would be sufficient.

6. Place recipient mice into a cage and supply them with food and acidified water containing neomycin. To prevent development of opportunistic infections in mice, keep the mice on acidified water for 6 weeks. Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

7. Six weeks after transplantation, warm the recipients, place them into a holder, and bleed 2 large drops into a 1.5-ml microcentrifuge tube filled with 1.0 ml of 200 mg/liter EDTA/PBS. 8. Centrifuge cells 3 min at 300 × g, room temperature. Add PharmLyse (or preferred red blood cell lysis buffer) according to the manufacturer’s instructions.

2A.6.12 Supplement 4

Current Protocols in Stem Cell Biology

Figure 2A.6.4 An example of how donor, recipient, and carrier cells can be distinguished following hematopoietic reconstitution. Dead cells and debris are excluded according to the uptake of 7-AAD and forward scatter profile; donor cells can subsequently be identified as Ly5.2/2 cells, recipient as Ly5.1/1, and carrier as Ly5.2/1.

9. Wash cells in 1.0 ml FACS wash buffer and centrifuge 5 min at 300 × g, 4◦ C. 10. Stain with 100 µl Ly5.1/Ly5.2 antibodies. Determine donor, recipient, and carrier cell populations using flow cytometry (see Fig. 2A.6.4 for an example).

PREPARATION OF ACIDIFIED DRINKING WATER FOR IRRADIATED MICE

SUPPORT PROTOCOL 3

The following acidic drinking water containing antibiotic should be provided to experimental mice for the first 6 weeks following irradiation to prevent opportunistic infection.

Materials Concentrated HCl Neomycin (Sigma) 1. Prepare a 100× stock HCl solution by adding 10 ml concentrated HCl to 830 ml water. 2. Prepare a 100× stock neomycin solution by adding 16.7 g neomycin to 100 ml water (keep in light-protected bottle up to 2 months at 4◦ C). 3. Prepare acidic drinking water by diluting 1 part of each stock solution in 100 parts of water before supplying it to mice.

ENDOTHELIAL ASSAY Using co-culture with the OP9 cell line, the ability of cells to differentiate towards the endothelial lineage and the capacity of endothelium to form networks can be tested. The method described here is adopted from the original technique described by Nishikawa et al. (1998) and Fraser et al. (2003).

BASIC PROTOCOL 7

Materials OP9 cells (see Support Protocol 4) Dissected E11.5 embryonic tissues Endothelial growth medium (see recipe) Anti-PECAM-1 antibody (see Support Protocol 5) 4-well tissue culture flat bottom plates (Nunc)

Somatic Stem Cells

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Supplement 4

Figure 2A.6.5 Endothelial tubule forming potential of flow cytometrically purified AGM region cell populations. (A) In vitro endothelial tubule forming potential of E11.5 AGM region cell populations is largely restricted to the VE-cadherin+ CD45− (endothelial) fraction. (B) Example of PECAM-1+ endothelial tubules produced from 5000 VE-cadherin+ CD45− cells after 4 days in culture. (C) Example of the extensive vascular networks produced from 20,000 VE-cadherin+ CD45− cells. Original magnification of photomicrographs 40×. VE-cad, VE-cadherin.

1. Grow OP9 cells as described in Support Protocol 4. 2. Prepare confluent layer of OP9 cells in multi-well plates. 3. From a single-cell suspension, isolate defined cellular populations from E11.5 AGM region by flow cytometry (see Basic Protocols 2 and 4), e.g., and plate them in endothelium growth medium on a confluent layer of OP9. Endothelial tubules have been produced from as few as 500 VE-cadherin+ CD45− cells (∼1500 cells/ml) from the E11.5 AGM region.

4. Assess endothelial tubule and network formation after 4 days of culture using antiPECAM-1 antibody staining (see Support Protocol 5). See Figure 2A.6.5 for the expected results of endothelial differentiation from E11.5 AGM cell populations purified according to the expression of VE-cadherin and CD45. SUPPORT PROTOCOL 4

MAINTENANCE OF OP9 CELLS This method describes how to maintain OP9 cells prior to co-culture.

Materials

Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

2A.6.14 Supplement 4

OP9 cells (developed by Nakano et al., 1994) Culture medium (see recipe), prewarmed CMF-PBS Dissociation solution (see recipe) 10% DMSO (BDH) 10-ml tube 75-cm2 tissue culture flasks (Iwaki) 10-ml plastic pipets 1-ml cryotubes (Nunc) 37◦ C, 5% CO2 incubator Current Protocols in Stem Cell Biology

1. Thaw an aliquot of cryopreserved OP9 cells (typically stored in 0.5 ml freezing medium) quickly at 37◦ C. 2. Transfer entire 0.5-ml volume of OP9 cells to a 10-ml tube containing 9.5 ml of prewarmed culture medium to dilute DMSO. 3. Centrifuge 3 min at 200 × g, room temperature. 4. Remove supernatant and resuspend the cell pellet in 10 ml prewarmed culture medium; transfer the cells into a 75-cm2 tissue culture flask. 5. Grow the cells to sub-confluency (no more than 80%), otherwise they become large vacuolated cells and irreversibly lose their essential properties, in a 37◦ C, 5% CO2 incubator. Typically, the cells will reach 80% confluency in 3 days.

6. To passage the cells, aspirate medium and add 10 ml of CMF-PBS. Repeat the procedure and aspirate PBS. 7. Add 2.0 ml dissociation solution to the cells and incubate 2 to 5 min at 37◦ C. Observe the dissociation under a microscope until single-cell suspension is obtained (∼2 to 5 min). 8. Add 8.0 ml prewarmed culture medium to neutralize trypsin. Collect and gently resuspend cells with a 10-ml plastic pipet; transfer to a 10-ml tube and centrifuge 3 min at 200 × g, room temperature. 9. For maintenance of OP9 cells in culture, remove supernatant, resuspend cells in 5 ml prewarned culture medium, and dispense into four new 75-cm2 flasks (1/4 of suspension per flask.) See Support Protocol 5 for preparation of OP9 cells for co-culture experiments. 10. Freeze 1 × 106 OP9 cells in 0.5 ml culture medium supplemented with 10% DMSO in 1-ml cryotubes by placing them first into a −80◦ C freezer and on the following day (or later) into liquid nitrogen.

VISUALIZATION OF ENDOTHELIAL TUBULES To confirm the presence of endothelial development, it is necessary to stain the product of co-cultures with antibodies specific for endothelium-associated antigens. A method for the rapid immunohistochemical visualization of PECAM-1 expression is described here.

SUPPORT PROTOCOL 5

NOTE: The method described is for the staining of cells co-cultured in a 4-well plate (Nunc).

Materials Cultures of endothelial cells (OP9 stroma; Support Protocol 4) in 4-well plates (Nunc) CMF-PBS 2% (w/v) paraformaldehyde(PFA)/PBS, pH 7.4 (Sigma) 0.1% (v/v) Nonidet P40 (NP40)/PBS (Sigma) 10% FBS/PBS Anti-PECAM-1 antibody (BD Bioscience) Secondary anti-rat IgG antibody conjugated with alkaline phosphatase (AP) (Southern Biotechnology Associates) 0.1 M Tris·Cl, pH 8.2 0.125 M Levamisol (Vector)

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Vector Blue alkaline phosphatase substrate kit III (Vector) 4-well tissue culture flat-bottom plates (Nunc) Microscope with camera attached 1. To prepare cells for co-culture experiments, prepare OP9 cells as described in Support Protocol 4. From the 5-ml single-cell suspension of OP9 cells, add a 0.5-ml aliquot to each well of a 4-well plate (this will be sufficient to generate a confluent stromal layer within 1 to 2 days). 2. Carefully remove culture medium from the wells of endothelial cell culture in a 4-well plate. 3. Wash two times with 500 µl CMF-PBS. Before removal of the last portion of PBS, tilt the plate carefully. 4. Add 500 µl of 2% PFA/PBS and incubate 20 min at room temperature. 5. Wash two times with 500 µl CMF-PBS. 6. Add 500 µl of 0.1% NP40/PBS and incubate 10 min at room temperature. 7. Wash two times with 500 µl CMF-PBS. 8. Block with 500 µl of 10% FBS/PBS 30 min at room temperature. 9. Remove blocking solution (10% FBS/PBS) and add the anti-PECAM-1 antibody (5 µg/ml) in 250 µl of 5% FBS/PBS. Incubate 1 hr at room temperature. 10. Wash three times with 500 µl CMF-PBS. 11. Add anti-rat IgG-AP (1:250) in 250 µl of 5% FBS/PBS and incubate 1 hr at room temperature. 12. Wash three times with 500 µl CMF-PBS. 13. According to the manufacturer’s instructions, add 0.1 M Tris·Cl, pH 8.2, plus 0.125 M Levamisol and incubate 15 min at room temperature. 14. Perform alkaline phosphatase staining using the Vector Blue AP substrate kit III according to the manufacturer’s instructions. 15. Wash two times with 500 µl CMF-PBS 16. Replace with distilled water. 17. Take photographs under a microscope. Store up to 1 month at 4◦ C if required. Figures 2A.6.5B and 2A.6.5C show anti-PECAM1 immunostained endothelial cultures.

MESENCHYMAL LINEAGE DIFFERENTIATION ASSAYS

Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

Mesenchymal stem and/or progenitor cells derived from embryonic tissues such as the AGM can be identified by well-defined in vitro differentiation assays. When cultured under the appropriate conditions, these cells are able to form tissue aggregates with characteristics that may resemble tissues such as bone, fat, and cartilage. Widely associated with osteogenic differentiation is the formation of colonies that express alkaline phosphatase, an early marker for osteoblasts, and later display mineralized nodules. Adipocytes are differentiated fat cells with very distinct morphology due to large lipid deposits. Cartilaginous tissue is composed of abundant extracellular matrix rich in proteoglycans in which chondrocytes are embedded.

2A.6.16 Supplement 4

Current Protocols in Stem Cell Biology

Osteogenic Differentiation of Mesenchymal Cells Primary cells can be cultured under different conditions to test for osteoblastic potential.

BASIC PROTOCOL 8

Materials Primary cell suspension (see Basic Protocol 2) Osteogenic differentiation medium (see recipe) DPBS 4% (w/v) paraformaldehyde (PFA) in PBS Alkaline phosphatase staining kit (Sigma Diagnostics cat. no. 85L1) or alizarin red (see Support Protocol 6) 6-well plates 37◦ C, 5% CO2 humidified incubator 1. Isolate primary cells and prepare a single-cell suspension (see Basic Protocol 2). 2. Prepare 6-well plates with 3 ml/well osteogenic differentiation medium (4◦ C). 3. Add cells at the densities specified in Table 2A.6.3, and culture for 10 to 12 days (for osteogenic potential) or 21 days (for mineralized colonies) in a 37◦ C, 5% CO2 humidified incubator. 4. On desired assay date, wash plates two times with 3 ml DPBS. 5. Fix cells with 2 ml of 4% PFA for 15 min at room temperature and wash two times with 3 ml distilled water. Table 2A.6.3 Cell Seeding Densities for Mesenchymal Differentiation Assays

Osteogenic assay (cells/cm2 )

Adipogenic assay (cells/cm2 )

Chondrogenic assay (cells/pellet)

5 × 103 to 5 × 104

5 × 103 to 5 × 104

5 × 105 to 5 × 106

Aorta-gonadmesonephros (AGM)

1 × 103

5 × 103

1 × 105

Liver

1 × 104

1 × 104

1 × 106

Midgestation tissue Yolk sac

Figure 2A.6.6 Mesenchymal cells from midgestation hematopoietic tissues. Differentiation to (A) osteogenic, (B) adipogenic, and (C) chondrogenic lineages. After 10 to 12 days in osteogenic medium, colonies of cells are positive (blue) when stained for alkaline phosphatase activity. After stimulation in adipogenic medium, colonies contained cells with a distinct adipocyte morphology, which includes the lipid droplets. After 21 days in chondrogenic medium, cells formed a cartilage-like tissue with an extracellular matrix rich in proteoglycans, as detected by toluidine blue staining. Somatic Stem Cells

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6a. For 10- to 12-day cultures, stain plates with alkaline phosphatase following the manufacturer’s instructions to visualize colonies with osteoblastic potential (see Fig. 2A.6.6A). 6b. For 21-day cultures, stain plates with alizarin red to visualize mineralized colonies (see Support Protocol 6). ALTERNATE PROTOCOL 1

Adipogenic Differentiation of Mesenchymal Cells Primary cells are cultured under different conditions to test for adipogenic potential and differentiation.

Additional Materials (also see Basic Protocol 8) Adipogenic differentiation medium I (see recipe) Adipogenic differentiation medium II (see recipe) 1. Isolate primary cells and prepare a single-cell suspension (see Basic Protocol 2). 2. Prepare 6-well plates with 3 ml/well adipogenic differentiation medium I. 3. Add cells at the densities specified in Table 2A.6.3, and culture for 2 to 3 days in a 37◦ C, 5% CO2 humidified incubator. 4. Remove adipocyte differentiation medium I and replace with 3 ml/well adipogenic differentiation medium II. 5. Culture cells for an additional 7 to 10 days in a 37◦ C, 5% CO2 humidified incubator. 6. Wash plates two times in 3 ml DPBS and analyze microscopically for cells with adipocyte morphology (i.e., lipid vacuoles; Fig. 2A.6.6B). 7. Occasionally, fix cells with 3 ml of 4% paraformaldehyde 15 min at room temperature. Wash two times with 3 ml DPBS. 8. Stain with oil red O (see Support Protocol 7), which stains lipoproteins red. ALTERNATE PROTOCOL 2

Chondrogenic Differentiation of Mesenchymal Cells Primary cells can be cultured under conditions that lead to chondrogenic differentiation.

Additional Materials (also see Basic Protocol 8) Chondrogenic differentiation medium (see recipe) Toluidine blue stain (see Support Protocol 8) Tissue tek 15-ml polypropylene tubes Plastic molds Cryostat 1. Isolate primary cells and prepare a single-cell suspension (see Basic Protocol 2). 2. Place cells in a 15-ml polypropylene tube and centrifuge 5 min at 1000 × g, 4◦ C, to form a micro-mass (Dennis et al., 1999 and Table 2A.6.3). Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

3. Culture the cell micro-mass (or aggregate) in 1 ml chondrogenic differentiation medium in 15-ml polypropylene tubes for 21 days in a 37◦ C, 5% CO2 humidified incubator (see cell numbers per tube in Table 2A.6.3). 4. Remove medium and wash cell micro-masses in 3 to 5 ml DPBS.

2A.6.18 Supplement 4

Current Protocols in Stem Cell Biology

5. Embed cell aggregates in tissue tek in a plastic mold. Then, very quickly freeze by placing the mold in a mixture of dry ice and 100% ethanol. As soon as the tissue tek solidifies, store the sample at −80◦ C. Prepare cryosections 8- to 10-µm thick on a cryostat. 6. Perform toluidine blue staining on sections to reveal proteoglycans in the extracellular matrix of the chondrogenic tissue (see Support Protocol 8). 7. Immunostain for collagen type II on sections of cell aggregates to confirm the cartilaginous nature of the cultured tissue (see Support Protocol 9). See Figure 2A.6.6 for the expected results of osteogenic (A), adipogenic (B), and chondrogenic (C) differentiation from E11 AGM cells.

HISTOLOGICAL STAINING WITH ALIZARIN RED FOR IDENTIFICATION OF BONE TISSUE

SUPPORT PROTOCOL 6

To confirm the presence of differentiated mesenchymal cells, it is necessary to stain the cells after culture with dyes specific for bone, fat, or cartilage-related molecules: respectively, alizarin red stains calcium deposits; oil red O stains lipid vacuoles; and toluidine blue stains proteoglycans. A method for the rapid histochemical verification of lineage differentiation is described here.

Materials Alizarin red (Sigma cat. no. A5533) 1 M NaOH Cultures of cell to be tested for differentiation PBS 4% (w/v) paraformaldehyde (PFA) 45-µm filter 1. Prepare a 0.2% (w/v) alizarin red solution in distilled water, adjust the pH to 4.2 with 1 M NaOH and filter using a 45-µm fitter. 2. Wash the cultures two times in 3 ml PBS by adding PBS on top of the cell monolayer and then removing it. 3. Fix cultures in 2 ml of 4% PFA 15 min at room temperature. 4. Wash the fixed cultures two times with 2 ml distilled water. 5. Add to each well, 2 to 3 ml of 0.2% alizarin red solution and stain for up to 10 min at room temperature. 6. Remove the staining solution and wash three times with 3 ml distilled water. 7. Visualize the mineralized tissue under a microscope. If mineralized tissue nodules have been formed, the calcium ions will stain bright red.

HISTOLOGICAL STAINING WITH OIL RED O STAIN FOR IDENTIFICATION OF ADIPOCYTES

SUPPORT PROTOCOL 7

Oil red O stains lipid vacuoles in cells identifying them as adipocytes.

Materials Oil red O (Sigma cat. no. 75087) Cultures of cells to be tested for differentiation in 6-well plates PBS 4% (w/v) paraformaldehyde (PFA)

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1. Prepare a 0.5% oil red O solution following the manufacturer’s instructions. 2. Wash the cultures two times in 3 ml PBS by adding PBS on top of the cell monolayer and then removing it. 3. Fix in 2 ml of 4% PFA 15 min at room temperature. 4. Wash the fixed cultures two times with 3 ml distilled water. 5. Add 2 to 3 ml of 0.5% oil red O solution to the wells and stain for up to 30 min at room temperature. 6. Remove the staining solution and wash three times with 3 ml distilled water. 7. Visualize the lipid deposits that are stained red. SUPPORT PROTOCOL 8

HISTOLOGICAL STAINING WITH TOLUIDINE BLUE STAIN FOR IDENTIFICATION OF CARTILAGE Cryostat sections of micro-mass cultures are stained with toluidine blue to detect proteoglycans, which are found in cartilage.

Materials Toluidine blue (Sigma cat. no. 89640) Cryostat sections of micro-mass cultures 4% paraformaldehyde (PFA) 45-µm filters 1. Prepare a 0.1% (w/v) toluidine blue solution in distilled water and filter using a 45-µm filter. 2. Fix the cryo-sections by submerging in 4% PFA for 5 min at room temperature. 3. Wash the fixed sections thoroughly with 150 to 200 ml distilled water. 4. Stain slides by submerging in 0.1% toluidine blue solution for 1 to 2 min. 5. Wash with 150 to 200 ml distilled water. 6. Visualize both the morphology and proteoglycan content of the tissue under the microscope. Cartilage tissue is composed of abundant extracellular matrix composed of proteoglycans that the toluidine solution stains purple. Morphologically, chondrocytes can also be detected embedded in this extracellular matrix (see Fig. 2A.6.6C) SUPPORT PROTOCOL 9

IMMUNOSTAINING SECTIONS OF MICRO-MASS CULTURES WITH ANTI-COLLAGEN TYPE II FOR IDENTIFICATON OF CARTILAGE To confirm the presence of differentiated cartilage tissue, it is necessary to stain the tissue after culture with antibody specific for collagen type II. This collagen forms the major part of the extracellular matrix that defines this tissue. A method for rapid immunostaining following chondrogenic differentiation is described here.

Materials Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

2A.6.20 Supplement 4

Cryosections of micro-mass cultures 4% (w/v) PFA PBS/0.05% (v/v) Tween 20 Collagen type II antibody (CIIC1, Developmental Studies Hybridoma Bank) Anti-immunoglobulin-HRP (Dako) Chromogen diaminobenzidine (DAB, Dako) Current Protocols in Stem Cell Biology

1. Fix the cryosections by submerging in 100 to 200 µl of 4% PFA per cryosection. 2. Wash the fixed sections two times with 150 to 200 ml distilled water. 3. Submerge the sections for 5 min in 150 to 200 ml PBS/Tween 20. 4. Cover the tissue sections with 100 to 200 µl collagen type II–specific antibody for 30 min. 5. Wash with 100 to 200 ml of PBS/Tween 20. 6. Incubate with 100 to 200 µl/cryosection secondary antibody, anti-mouse immunoglobulin-HRP 30 min and wash abundantly with PBS/Tween 20. 7. Cover the sections with 100 to 200 µl DAB substrate for 3 min. 8. Wash with 100 to 200 ml distilled water. 9. Visualize under the microscope. Tissue expressing collagen type II will stain brown. No brown stain should be detected if collagen type II is not present.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Adipogenic differentiation medium I Prepare a solution of DMEM containing with 10% FBS and 100 U penicillin/100 mg streptomycin per 500 ml medium. Store up to 1 month at 4◦ C.

Adipogenic differentiation medium II DMEM containing: 1% FBS 100 U penicillin/100 mg streptomycin per 500 ml medium 10−7 M dexamethasone (Sigma cat. no. D8893) 100 ng/ml insulin (Sigma cat. no. I0516) Store up to 1 month at 4◦ C Chondrogenic differentiation medium DMEM containing: Insulin-transferrin-selenium (ITS+ ; Sigma cat. no. I2521) 100 U penicillin/100 mg streptomycin per 500 ml medium 0.1 mM L-ascorbic acid 2-phosphate 10−9 M dexamethasone 20 ng/ml TGF-1 (RD Systems cat. no. 240-B-002) Prepare fresh Collagenase type I For a 20× collagenase type I (Sigma) stock solution: prepare a 2.5% collagenase type I stock solution in medium II (see recipe). Keep at −20◦ C until needed.

Culture medium αMEM (Invitrogen) supplemented with: FBS (20%) (Invitrogen) Glutamine (4 mM) (Invitrogen) continued

Somatic Stem Cells

2A.6.21 Current Protocols in Stem Cell Biology

Supplement 4

2-Mercaptoethanol (0.1 mM) Penicillin (50 U/ml) Streptomycin (50 µg/ml) Store up to 2 months at −20◦ C Dissociation solution Trypsin (0.025%; Invitrogen) Chicken serum (0.1%; Flow Labs) EDTA (1.3 mM; Sigma) Store up to 2 months at −20◦ C Endothelial growth medium αMEM medium containing: 10% FBS 4 mM glutamine 0.1 mM 2-mercaptoethanol 50 U/ml penicillin 50 µg/ml streptomycin 50 ng/ml vascular endothelial growth factor (VEGF; PeproTech)

Medium I (medium for collection of embryos) Dulbecco’s phosphate buffered saline (PBS) with Ca2+ and Mg2+ containing: Penicillin (100 U/ml) Streptomycin (100 µg/ml) Store up to 2 months at −20◦ C Medium II (medium for dissections) PBS with Ca2+ and Mg2+ (Sigma) 7% fetal bovine serum (FBS; Invitrogen) Penicillin (100 U/ml) Streptomycin (100 µg/ml) Medium III Dulbecco’s PBS (DPBS; Ca2+ /Mg2+ -free). Store at room temperature.

Millipore Durapore membrane filters Before use, wash and sterilize Millipore Durapore 0.65-µm membrane filters in several changes of boiling tissue culture water (Sigma cat. no. W-3500). Store at room temperature.

Osteogenic differentiation medium Dulbecco’s modified Eagle’s medium (DMEM) containing: 15% FBS 100 U penicillin/100 mg/ml streptomycin per 500 ml 0.2 mM L-ascorbic acid 2-phosphate (Sigma cat. no. A8960) 0.01 M glycerophosphate (Sigma cat. no. G9891) Store up to 1 month at 4◦ C Stainless steel wire mesh supports Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

Prepare stainless steel wire mesh supports by bending a 22 × 12–mm rectangular piece of mesh (5-mm height and 12 × 12–mm platform). Wash supports in HNO3 for 2 to 24 hr. Rinse five times in sterile Milli-Q water and then in 70% ethanol. Rinse two times in tissue culture water (Sigma cat. no. W-3500). Dry the supports in a tissue culture hood. Store at room temperature.

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COMMENTARY Background Information The hematopoietic system is an essential tissue system that provides for all blood cells in circulation and associated organs. It contains a variety of cells such as erythrocytes (necessary for oxygen transport), macrophages and natural killer cells (necessary for innate immunity), B and T lymphocytes (necessary for acquired immunity), and other differentiated cell types providing unique functions. In adult mammals, it is found in hematopoietic stem cells that reside in the bone marrow. While much is known concerning the differentiation of blood cells in the adult, there is intense current interest in the embryonic origins of the adult hematopoietic system and particularly hematopoietic stem cells (Jaffredo et al., 2005; Tavian and Peault, 2005). The hematopoietic system in mammalian embryos develops in a spatial and temporal association with the vasculature. The earliest origins of the hematopoietic system in the mouse have been mapped and quantified in the intraembryonic [aorta-gonadmesonephros (AGM) and liver], extraembryonic tissues (yolk sac, placenta), and the blood vessels that link these two parts of the embryo (vitelline and placenta vessels; Moore and Metcalf, 1970; Muller et al., 1994; Medvinsky and Dzierzak, 1996; de Bruijn et al., 2000; Cumano et al., 2001; Kumaravelu et al., 2002; Gekas et al., 2005; Ottersbach and Dzierzak, 2005). Various types of hematopoietic progenitors (mature and immature), multipotential progenitors, and hematopoietic stem cells (neonatal and adult repopulating) have been described (Moore and Metcalf, 1970; Cumano et al., 1993; Medvinsky et al., 1993; Yoder et al., 1995). Yet there is need for further dissection, characterization, and manipulation of these (and perhaps other) early hematopoietic cell types to resolve on-going controversies of how the system is first generated, subsequently expanded, and maintained. The potency and function of hematopoietic cells produced by the yolk sac and placenta as compared to those produced by the intra-body portion of the mouse embryo is currently under investigation and continues to pose new questions and research in the field of developmental hematopoiesis (Robin et al., 2006). Also, the search for the direct precursor cells to hematopoietic cells continues and has focused

on the relationship of embryonic endothelial and mesenchymal cells as a potential source (de Bruijn et al., 2002; North et al., 2002; Bertrand et al., 2005). Pluripotential embryonic stem (ES) cells are a challenging additional source for the generation of hematopoietic stem cell precursors (Kennedy and Keller, 2003; Kennedy et al., 2007). Overall, the further understanding of the in vivo molecular and cellular interactions necessary for hematopoietic stem cell generation in the mammalian embryo offers great promise for the production of specific lineages and/or an entire adult hematopoietic system for clinical cell replacement therapies. The protocols presented here are designed to instruct fundamental research scientists in the dissection, preparation, culturing, and assaying of the first hematopoietic progenitors and stem cells as they appear in the mouse conceptus. Basic Protocol 1 describes the dissection of mouse embryonic tissues; Basic Protocol 2 describes the preparation of cell suspensions from midgestation mouse embryos; Basic Protocol 3 describes the culture of embryonic tissue explants, cells and cocultures used to support hematopoietic stem cells ex vivo; Basic Protocol 4 delineates the procedure for flow cytometric analysis; and for delineating the potential of embryonic cells Basic Protocols 6, 7, and 8 describe differentiation assays for hematopoietic, endothelial, and mesenchymal lineages, respectively. Basic Protocol 5 describes a long-term in vivo repopulation assay for stem cells.

Critical Parameters Most procedures described here require tissue culture facilities where cultures can be established and maintained under aseptic/sterile conditions—including flow hoods, incubators, and autoclaves. The incubations described should be performed in 37◦ C, 5% CO2 humidified incubators, unless otherwise specified. Production of embryos may undergo variations during the year. A cause of these variations has yet to be established. Periods of poor embryo production end at some point and are changed to good production periods without obvious reason. All dissections should be performed in solutions based on PBS with calcium and magnesium at room temperature and cell Somatic Stem Cells

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suspensions should be kept on ice in solutions based on PBS without calcium and magnesium. Staging of embryos is very important. E10 embryos below 36 sp are not capable of generating long-term repopulating HSCs in explant cultures. Preliminary work on familiarizing oneself with embryos of appropriate age, which includes counting somites and grouping of embryos, is required before performing actual experiments. Dissecting embryos requires experience. For the beginner, isolation of one AGM region from the embryo can take up to 20 to 30 min and still result in significantly damaged tissues. Systematic practice over 2 weeks is normally sufficient to reach good productivity. The experienced researcher is capable of dissecting ∼30 AGMs in 1 hr. While isolating the yolk sac, make sure that all large vessels connecting it with the body of the embryo are removed. Collagenase/dispase solutions are not very standard reagents and enzymatic activity may vary. Therefore, try various concentrations of this enzyme mixture to obtain desirable cell suspensions. Normally, excessive digestion time does not yield more progenitors and stem cells but rather generates more dead cells. Exposing cells to greater centrifugal forces than those recommended may result in a high proportion of dead cells in the resultant pellet and the formation of cell clumps, which are resistant to dissociation by pipetting. Be careful with setting the temperature in the refrigerated centrifuge: setting it <4◦ C may give temperatures below 0◦ C due to temperature fluctuations in the centrifuge. Normal experiments will be with relatively small numbers of cells, make sure that when supernatants are removed that a small amount of medium (between 20 and 50 µl) is left above the pellet. The CFU-C and HSC content may vary depending on different strains of mice. While growing OP9 cells, make sure that they do not reach 100% confluency, as their properties may irreversibly change.

Anticipated Results

Analysis and Manipulation of Hematopoietic Progenitor and Stem Cells from Murine Embryonic Tissues

From one freshly prepared E11.5 AGM region, it is expected that ∼1 HSC and 122 ± 32 CFU-Cs can be detected; HSCs will be enriched within the VE-cadherin+ CD45+ population while the majority of CFU-Cs will be present in the VE-cadherin− CD45+ population. With regard to the mesenchymal lineages, it is expected that 81 ± 10 osteogenic,

124 ± 32 adipogenic, and 28 ± 1 chondrogenic progenitors can be detected. The vast majority of in vitro endothelial differentiation potential will be found within the VEcadherin+ CD45− population. From E11.5 AGM regions developed by the explant culture technique, it is expected that 10 to 12 HSCs and 147 ± 21 CFU-Cs will be present within a single organ.

Time Considerations It is not recommended to carry out dissections for longer than 3 hr. Enzymatic digestion and preparation of cell suspensions take ∼1.5 hr; staining with antibodies requires 1 hr; cell sorting requires 1 hr; preparations of aliquots and setting up methylcellulose cultures require1 hr. It takes between 8 and 10 days to obtain results of colony growth in methylcellulosebased medium and 4 days for endothelial tube formation. For transplantation experiments, preparation of recipient mice (including irradiation) may overlap with preparation of cell suspensions; therefore, plan the experiment carefully and seek assistance if necessary. The duration of these experiments is >3 months.

Literature Cited Bertrand, J.Y., Giroux, S., Golub, R., Klaine, M., Jalil, A., Boucontet, L., Godin, I., and Cumano, A. 2005. Characterization of purified intraembryonic hematopoietic stem cells as a tool to define their site of origin. Proc. Natl. Acad. Sci. U.S.A. 102:134-139. Cumano, A., Furlonger, C., and Paige, C.J. 1993. Differentiation and characterization of B-cell precursors detected in the yolk sac and embryo body of embryos beginning at the 10- to 12-somite stage. Proc. Natl. Acad. Sci. U.S.A. 90:6429-6433. Cumano, A., Ferraz, J.C., Klaine, M., Di Santo, J.P., and Godin, I. 2001. Intraembryonic, but not yolk sac hematopoietic precursors, isolated before circulation, provide long-term multilineage reconstitution. Immunity 15:477-485. de Bruijn, M.R.T.R., Speck, N.A., Peeters, M.C.E., and Dzierzak, E. 2000. Definitive hematopoietic stem cells first emerge from the major arterial regions of the mouse embryo. EMBO J. 19:2465-2474. de Bruijn, M., Ma, X., Robin, C., Ottersbach, K., Sanchez, M.-J., and Dzierzak, E. 2002. HSCs localize to the endothelial layer in the midgestation mouse aorta. Immunity 16:673-683. Dennis, J.E., Merriam, A., Awadallah, A., Yoo, J.U., Johnstone, B., and Caplan, A.I. 1999. A quadripotential mesenchymal progenitor cell isolated from the marrow of an adult mouse. J. Bone Miner. Res. 14:700-709.

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Fraser, S.T., Ogawa, M., Yokomizo, T., Ito, Y., and Nishikawa, S. 2003. Putative intermediate precursor between hematogenic endothelial cells and blood cells in the developing embryo. Dev. Growth Differ. 45:63-75. Gekas, C., Dieterlen-Lievre, F., Orkin, S.H., and Mikkola, H.K. 2005. The placenta is a niche for hematopoietic stem cells. Dev. Cell 8:365-375. Hogan, B.C.F. and Beddington R. 2002. Manipulating the Mouse Embryo. Cold Spring Harbor Laboratory Press. Woodbury, NY. Jaffredo, T., Nottingham, W., Liddiard, K., Bollerot, K., Pouget, C., and de Bruijn, M. 2005. From hemangioblast to hematopoietic stem cell: An endothelial connection? Exp. Hematol. 33:10291040. Kennedy, M. and Keller, G.M. 2003. Hematopoietic commitment of ES cells in culture. Methods Enzymol. 365:39-59. Kennedy, M., D’Souza, S.L., Lynch-Kattman, M., Schwantz, S., and Keller, G. 2007. Development of the hemangioblast defines the onset of hematopoiesis in human ES cell differentiation cultures. Blood 109:2679-2687. Kumaravelu, P., Hook, L., Morrison, A.M., Ure, J., Zhao, S., Zuyev, S., Ansell, J., and Medvinsky, A. 2002. Quantitative developmental anatomy of definitive haematopoietic stem cells/longterm repopulating units (HSC/RUs): Role of the aorta-gonad- mesonephros (AGM) region and the yolk sac in colonisation of the mouse embryonic liver. Development 129:48914899. Medvinsky, A. and Dzierzak, E. 1996. Definitive hematopoiesis is autonomously initiated by the AGM region. Cell 86:897-906. Medvinsky, A.L., Samoylina, N.L., Muller, A.M., and Dzierzak, E.A. 1993. An early pre-liver intraembryonic source of CFU-S in the developing mouse. Nature 364:64-67. Moore, M.A. and Metcalf, D. 1970. Ontogeny of the haemopoietic system: Yolk sac origin of in vivo and in vitro colony forming cells in the developing mouse embryo. Br. J. Haematol. 18:279296.

Muller, A.M., Medvinsky, A., Strouboulis, J., Grosveld, F., and Dzierzak, E. 1994. Development of hematopoietic stem cell activity in the mouse embryo. Immunity 1:291-301. Nakano, T., Kodama, H., and Honjo, T. 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265:10981101. Nishikawa, S.I., Nishikawa, S., Kawamoto, H., Yoshida, H., Kizumoto, M., Kataoka, H., and Katsura, Y. 1998. In vitro generation of lymphohematopoietic cells from endothelial cells purified from murine embryos. Immunity 8:761769. North, T.E., de Bruijn, M.F., Stacy, T., Talebian, L., Lind, E., Robin, C., Binder, M., Dzierzak, E., and Speck, N.A. 2002. Runx1 expression marks long-term repopulating hematopoietic stem cells in the midgestation mouse embryo. Immunity 16:661-672. Ottersbach, K. and Dzierzak, E. 2005. The murine placenta contains hematopoietic stem cells within the vascular labyrinth region. Dev. Cell 8:377-387. Robin, C., Ottersbach, K., Durand, C., Peeters, M., Vanes, L., Tybulewicz, V., and Dzierzak, E. 2006. An unexpected role for IL-3 in the embryonic development of hematopoietic stem cells. Dev. Cell 11:171-180. Taoudi, S., Morrison, A.M., Inoue, H., Gribi, R., Ure, J., and Medvinsky, A. 2005. Progressive divergence of definitive haematopoietic stem cells from the endothelial compartment does not depend on contact with the foetal liver. Development 132:4179-4191. Tavian, M. and Peault, B. 2005. The changing cellular environments of hematopoiesis in human development in utero. Exp. Hematol. 33:10621069. Yoder, M.C., King, B., Hiatt, K., and Williams, D.A. 1995. Murine embryonic yolk sac cells promote in vitro proliferation of bone marrow high proliferative potential colony-forming cells. Blood 86:1322-1330.

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High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

UNIT 2A.7

Sanja Sekulovic,1, 2 Suzan Imren,1 and Keith Humphries1, 2 1

Terry Fox Laboratory, British Columbia Cancer Agency, Vancouver, British Columbia, Canada 2 University of British Columbia, Vancouver, British Columbia, Canada

ABSTRACT Development of strategies to extensively expand hematopoietic stem cells (HSCs) in vitro will be a major factor in enhancing the success of a range of transplant-based therapies for malignant and genetic disorders. In addition to potential clinical applications, the ability to increase the number of HSCs in culture will facilitate investigations into the mechanisms underlying self-renewal. In this unit, we describe a robust strategy for consistently achieving over 1000-fold net expansion of HSCs in short-term in vitro culture by using novel engineered fusions of the N-terminal domain of nucleoporin 98 (NUP98) and the homeodomain of the hox transcription factor, HOXA10 (so called NUP98-HOXA10hd fusion). We also provide a detailed protocol for monitoring the magnitude of HSC expansion in culture by limiting dilution assay of competitive lymphomyeloid repopulating units (CRU Assay). These procedures provide new possibilities for achieving significant numbers of HSCs in culture, as well as for studying HSCs C 2008 biochemically and genetically. Curr. Protoc. Stem Cell Biol. 4:2A.7.1-2A.7.14.
by John Wiley & Sons, Inc. Keywords: NUP98-HOX fusion r HSC expansion r CRU assay r multilineage reconstitution

INTRODUCTION The establishment and subsequent lifelong maintenance of hematopoiesis relies on a rare subset of cells called hematopoietic stem cells (HSCs). HSCs are currently best defined based on their functional properties to self-renew, or divide in such a way that one or both of the progeny retain undiminished differentiation and proliferative potential, including the ability to produce progeny committed to differentiate along all of the hematopoietic lineages and to contribute to long-term lympho-myeloid hematopoiesis upon transplantation. The existence of HSCs with their capacity for sustained self-renewal and ability to re-establish long-term hematopoiesis is the basis of an increasing range of applications of HSC transplantation for the treatment of various malignant and genetic disorders (Shizuru et al., 2005; Verma and Weitzman, 2005). Broader use—e.g., from cord blood (CB) sources—and improved safety (e.g., by accelerating recovery) of such therapy would be greatly facilitated by the development of tools to achieve significant expansion of HSC numbers in vitro. In addition to potential clinical applications, the ability to extensively amplify the number of HSCs in culture will likely be instrumental in elucidating the complex and still poorly understood mechanisms underlying HSC behavior. Improved HSC purification techniques (Adolfsson et al., 2001; Christensen and Weissman, 2001; Chen et al., 2002; Uchida et al., 2003; Matsuzaki et al., 2004; Kiel et al., 2005; Wagers and Weissman, 2006; Balazs et al., 2006) and identification of extrinsic and intrinsic regulators of cell fate determination (see Background Information), as Somatic Stem Cells Current Protocols in Stem Cell Biology 2A.7.1-2A.7.14 Published online January 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a07s4 C 2008 John Wiley & Sons, Inc. Copyright


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well as functional HSC assays, have all contributed to systematic analysis of conditions that would support HSC maintenance or expansion in culture. Nevertheless, molecular and cellular signals that influence the choice between self-renewal and differentiation remain incompletely defined, thus increasing the challenge for expanding HSC populations in vitro. The protocols described in this chapter build on a recently developed strategy for consistently achieving over 1000-fold net expansion of HSCs in short-term (6- to 10-day) in vitro liquid culture by retroviral-mediated transfer of a novel engineered NUP98-HOX homeodomain fusion gene (NUP98-HOXA10hd) encoding a fusion protein consisting of the N-terminal domain of nucleoporin-98 (NUP98), which contains a region of multiple phenylalanine-glycine repeats that may act as transcriptional coactivator through binding to CBP/p300 (Kasper et al., 1999) and the 60-amino-acid DNA-binding domain (homeodomain) of HOXA10 (Ohta et al., 2007). Proviral integration analysis of BM DNA from recipients reconstituted with NUP98-HOXA10hd–transduced cells from cultures initiated with either large or limiting numbers of BM cells confirmed that NUP98-HOXA10hd fusion stimulates all transduced HSCs (rather than a minor subset of these cells) to expand in vitro (Ohta et al., 2007). Therefore, this strategy is easily adaptable to both polyclonal and clonal HSC expansion of transduced murine bone marrow cells maintained in static culture with defined cytokines for a relatively short period (10 days of total culture; 6 days post-transduction; see Basic Protocol 1). The estimation of the magnitude of HSC expansion achieved in culture is done by measuring the HSC frequency in initial versus culture containing NUP98-HOXA10HD-expanded HSCs, with a limiting dilution assay of long-term competitive lympho-myeloid repopulating units (CRU assay; see Basic Protocol 2). Description of these two protocols is shown in Figure 2A.7.1. Thus, the

Figure 2A.7.1 General experimental design for in vitro expansion of murine hematopoietic stem cells using NUP98HOXA10hd (NA10hd).

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procedure described allows, in 10 days of culture, the ready generation of >1000 clonally expanded HSC from a single initially transduced HSC or over a mouse equivalent of HSC (10,000) from as few as 10 starting HSC. Even higher levels of expansion appear feasible with modest extensions in the length of culture post transduction (>10,000-fold with 17 days culture; Ohta et al., 2007). The expansion procedure described thus provides a robust method for obtaining a population of primitive hematopoietic stem cells in vitro (defined by rigorous functional and quantitative assay of competitive lymphomyeloid repopulating unit frequency) strongly biased towards symmetrical self-renewal decisions and for obtaining large numbers of either clonally derived or polyclonal HSCs for subsequent in vitro or in vivo studies.

EX VIVO EXPANSION OF MURINE HSCs IN SHORT-TERM CULTURES Stable integration of murine retroviral vectors requires cell division of the target cells. Therefore, in order to activate quiescent HSCs into cell cycling, BM donor mice are injected with 5-fluorouracil (5-FU). This procedure removes a large proportion of actively cycling, more differentiated cells, thus increasing the frequency and triggering the cycling of HSCs that become more susceptible to retroviral infection (Harrison and Lerner, 1991; Bodine et al., 1991).

BASIC PROTOCOL 1

The protocol consists of a 2-day prestimulation period involving exposure to a combination of cytokines—interleukin-3 (IL-3), interleukin-6 (IL-6), and stem cell factor (SCF)—critical to maintain/trigger cycling and to promote survival of HSCs prior to and during the infection (Luskey et al., 1992; Bodine et al., 1989); this is followed by a 2-day infection period based on a well established method for achieving stable integration of a transgene with high efficiency, using recombinant murine retroviruses and a static liquid culture period for 6 days (or greater), allowing for post-infection expansion of transduced HSCs. A cytokine cocktail containing IL-3, IL-6, and SCF is added to the medium during the entire culture period. While this cytokine cocktail has proven sufficient for robust and high-level expansion of NUP98-HOXA10hd–transduced HSCs, it is likely that further optimization in the nature and concentration of growth factors is possible, to both reduce the output of differentiated cells in culture and increase the yield of HSCs. The polyclonal nature of the HSC expansion in cultures of bulk NUP98-HOXA10hdtransduced BM cells (Ohta et al., 2007) suggests a general susceptibility of HSCs to the effects of this fusion gene and provides a strategy to obtain large numbers of a complex, polyclonal population of expanded HSC. Alternatively, cultures can be initiated with reduced numbers of input cells (minicultures) containing an estimated 1 to 2 HSCs (either from unseparated 5-FU-pretreated BM or after isolation of the Sca1+ lin− or ckit+ Sca1+ lin− cells). Such an experimental design enables direct assessment of HSC expansion levels in the culture, as well as production of a clonally expanded HSC population that may be more suitable for certain applications. Although not described in detail below, preliminary experiments initiated with single CD45mid lin− Rholow SP (sidepopulation) cells (Uchida et al., 2003), followed by NUP98-HOXA10hd transduction, indicate the feasibility of achieving high-level HSC expansion from a single starting HSC, thus opening up additional avenues for rigorous examination of the clonal expansion and differentiation behavior of individual starting HSC.

Materials 2- to 4-month-old C57Bl/6Ly-Pep3b [Pep3b (Ly5.1)] mice, bred and maintained at the British Columbia Cancer Research Centre (http://www.bccrc.ca) animal facility according to the guidelines of the Canadian Council on Animal Care 5-fluorouracil (5-FU, Mayne Pharma, http://www.maynepharma.com/)

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Dulbecco’s phosphate-buffered saline without Ca2+ and Mg2+ (CMF-DPBS, StemCell Technologies, cat. no. 37350) CMF-PBS containing 2% (v/v) fetal bovine serum (FBS, StemCell Technologies, cat. no. 06250); store up to 1 month at 4◦ C 0.8% (w/v) NH4 Cl/1 mM EDTA in H2 O (StemCell Technologies, cat. no. 07850), ice cold DMEM with 15% FBS (see recipe) containing 10 ng/ml human IL-6, 6 ng/ml murine IL-3, and 100 ng/ml murine SCF (cytokines available from StemCell Technologies) Sca1+ lin− or c-kit+ Sca1+ lin− BM cells (to initiate culture with a starting population highly enriched in HSCs, 5-FU BM can be further purified to obtain these cells; see Ohta et al., 2007) GP+ E-86 retroviral producer cells (Dr. Keith Humphries, Terry Fox Laboratory, BC Cancer Agency, Vancouver, Canada; see annotation to step 7) irradiated with 40 Gy of X rays (or equivalent) DMEM with 15% FBS (see recipe) containing 10 ng/ml human IL-6, 6 ng/ml murine IL-3, 100 ng/ml murine SCF (StemCell Technologies), and 5 µg/ml protamine sulfate (Sigma) DMEM wash medium (see recipe) DMEM with 15% FBS (see recipe) Dissecting instruments including scissors and forceps 22-G, 1-in. and 26-G, 0.5-in. needles and 3-ml syringes for harvesting bone marrow Tabletop centrifuge Bacteriological petri dishes, standard style, 100 × 20–mm (BD Falcon, cat. no. 351005) 96-well U-bottom microtiter plates (BD Falcon, cat. no. 353077) Cell culture dishes, standard tissue culture treated, 100 × 20–mm (BD Falcon, cat. no. 353003) Cell lifters (Corning) 50-ml conical tubes (BD Falcon, cat. no. 352070) 24-well flat-bottom plates (BD Falcon, cat no. 353047) Additional reagents and equipment for intravenous injection of mice (Donovan and Brown, 2006a), euthanasia of mice (Donovan and Brown, 2006b), counting cells (Phelan, 2006), and CRU assay (Basic Protocol 2) NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Isolate, plate, and prestimulate murine BM cells 1. Day −4: Inject donor Pep3b (Ly5.1) mice intravenously in tail vein (Donovan and Brown, 2006a) with 150 mg/kg 5-FU dissolved in CMF-DPBS. Day 0: Harvest and plate BM cells and initiate prestimulation period (2 days) Day 0 of the protocol corresponds to “day 4” following the 5-FU injection in step 1.

High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

2. Sacrifice donor mice by CO2 asphyxiation (Donovan and Brown, 2006b). Harvest BM cells into 3 ml of CMF-DPBS with 2% FBS by flushing mouse femurs and tibias. Use a 3-ml syringe attached to a 22-G, 1-in. needle to flush femurs or a 26-G, 0.5-in. needle to flush tibias. 3. Lyse red blood cells by adding ∼10 ml ice-cold 0.8% NH4 Cl/0.1 mM EDTA and then incubate on ice for 5 to 10 min.

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4. Centrifuge cells 7 min at 300 × g, 4◦ C, remove the supernatant, resuspend pellet in 1 ml of DMEM with 15% FBS, and count an aliquot of cells (Phelan, 2006) using a 1:10 dilution. 5a. For bulk expansion (from nonlimiting numbers of starting HSC): Initiate individual cultures with 5-FU-pretreated BM cells (from step 4) at 3–5 × 105 cells/ml (estimated to contain a maximum of ∼60 to 100 HSCs assuming an HSC frequency of ∼1 in 5000 for day-4 5-FU BM; Ohta et al., 2007). Use DMEM supplemented with 15% FBS and cytokines (6 ng/ml of IL-3, 10 ng/ml of IL-6, 100 ng/ml of SCF) as a culture medium and plate cells in 100 × 20–mm bacteriological petri dishes to minimize adherence. Incubate cells. 5b. For expansion in “mini-cultures” (estimated to contain ∼1 to 2 HSC per culture): Initiate cultures with 5000 unseparated 5-FU-pretreated (from step 4) or 500 Sca1+ lin− or 30 c-kit+ Sca1+ lin− BM cells in 100 µl of DMEM supplemented with 15% FBS and cytokines (6 ng/ml of IL-3, 10 ng/ml of IL-6, 100 ng/ml of SCF) in individual wells of a 96-well U-bottom microtiter plate. Maintain the cell concentration of each “mini-culture” at 105 to 106 cells/ml (see day 7—step 14, below—for more details). Incubate cells. 6. Reserve a small amount of 5-FU-pretreated murine BM to perform the day-0 CRU assay (see Basic Protocol 2).

Infect BM cells by cocultivation Day 2: Initiate infection period (duration, 2 days) 7. In the morning, plate 6 × 106 irradiated GP+ E-86 retroviral producer cells in a 100 × 20–mm tissue culture–treated dish or 37,000 irradiated GP+ E-86 retroviral producer cells per well of a 96-well U-bottom plate (to achieve 90% confluence). As a control, we routinely use viral producers for GFP and, for HSC expansion, viral producers for NUP98-HOXA10hd – GFP (or NUP98-HOXB4 – GFP). All of our vectors are based on the murine stem cell virus (MSCV) internal ribosomal entry site (IRES) enhanced green fluorescent protein (GFP) (MSCV-IRES-GFP or GFP vector), which serves as a backbone for cloning of a NUP98-HOXA10hd and NUP98HOXB4 cDNA upstream of IRES to create MSCV-NUP98-HOXA10hd-IRES-GFP (NUP98-HOXA10hdGFP vector) and MSCV-NUP98-HOXB4-IRES-GFP (NUP98-HOXB4 vector). NUP98HOXA10hd and NUP98-HOXB4 vectors consist of a 409-amino-acid (exons 1 to 12) N-terminal region of nucleoporin-98 (NUP98) and the 60-amino-acid homeodomain of HOXA10 exon2 and homeobox-containing exon of HOXB4 respectively (previously described in Pineault et al., 2004) and are available upon request (Dr. Keith Humphries, Terry Fox Laboratory, BC Cancer Agency, Vancouver, Canada). Production of high-titer helper-free retrovirus was carried out by standard procedures, using virus-containing supernatants from transfected amphotropic Phoenix packaging cells to infect the ecotropic packaging cell line GP+ E86 (described in Kalberer et al., 2002).

8. In the afternoon, harvest BM cells (from steps 5a or 5b; end of prestimulation) by scraping the plates with a cell lifter (or by scraping the wells with a pipet tip). Centrifuge cells 7 min at 300 × g, room temperature. Remove the supernatant, count an aliquot of cells (Phelan, 2006), and resuspend the cells in 7 ml (or 100 µl for “mini-cultures” in wells of 96-well plate) of DMEM with 15% FBS supplemented with cytokines (6 ng/ml of IL-3, 10 ng/ml of IL-6, 100 ng/ml of SCF) and 5 µg/ml protamine sulfate. 9. Remove medium from viral producer plate(s) or wells (step 7) and gently (dropwise) add BM cells (resuspended in 7 ml for plate or 100 µl for well; see step 8) on top of irradiated producers. Incubate cells. Be sure not to place more than 5 × 106 BM cells into a 100 × 20–mm dish.

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End infection and initiate expansion period Day 4 10. Remove BM cells from adherent retroviral producer cells, being careful not to disrupt the producers. Recover as many BM cells as possible by gently (dropwise) washing the surface of the producer cells with 10 ml (or 200 µl for “mini-cultures”) of DMEM wash medium. Repeat the washing three to five times. 11. Combine all BM cells in a 50-ml conical tube, and pellet by centrifuging for 7 min at 300 × g, room temperature. 12. Replate in a 100 × 20–mm bacteriological petri dish at 5 × 105 cells/ml and culture using 10 ml DMEM with 15% FBS supplemented with cytokines (without protamine sulfate). Incubate cells.

Day 6 13. If using a FACS-selectable marker (e.g., GFP), determine gene transfer efficiency by flow cytometry (Robinson et al., 2007; see Anticipated Results for expected gene transfer rate). Day 7 14. Harvest BM cells in suspension and by scraping the plates with a cell lifter, count an aliquot (Phelan, 2006), resuspend cells in 10 ml fresh DMEM with 15% FBS supplemented with cytokines, and replate 10% of the initial culture into the same size dish (i.e., a 1:10 split to keep cell density to appropriate levels). Continue cultures until day 10. This provides enough cells for extensive quantification of HSC content by limiting dilution analysis and for phenotyping. Of course, if greater numbers are required (e.g., for proviral integration analysis by Southern blot hybridization) at the end of the culture, largervolume cultures can be set up at the time of the split. In the case of mini-cultures (initiated in 96-well plates), the whole culture is simply transferred at day 7 to individual wells of 24-well plate in 500 µl of fresh DMEM with 15% FBS supplemented with cytokines.

Day 10 15. Harvest BM cells in suspension and by scraping plates with cell lifter, count an aliquot (Phelan, 2006), resuspend cells in 10 ml fresh DMEM with 2% FBS (no cytokines), and prepare various desired doses of test cells for day-10 CRU assay (see Basic Protocol 2). BASIC PROTOCOL 2

High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

QUANTITATION OF MURINE HSCs BY LIMITING DILUTION ANALYSIS IN COMPETITIVELY REPOPULATED HOSTS Changes in HSC numbers are calculated from a comparison of the number of CRUs measured in the starting population of 5-FU-pretreated BM cells (day-0 CRU assay; also see Basic Protocol 1, step 6) versus the number of transduced GFP+ CRUs, measured at the end of the culture period (day-10 CRU assay; also see Basic Protocol 1, step 15). The CRU assay provides the specificity required for the exclusive quantification of hematopoietic stem cells with life-long blood cell–producing activity (Szilvassy et al., 1990). This procedure uses the principles of limiting dilution analysis to measure the frequency of cells in a given suspension that have transplantable long-term repopulating ability and can individually generate both lymphoid and myeloid progeny. Normal mice are pretreated with a lethal dose of radiation (myeloablative treatment), while c-kit mutant mice—whose stem cells are defective (Miller et al., 1996)—are treated with a sublethal dose. The treatment of the hosts maximizes the sensitivity of the assay and reduces the competing endogenous stem cell population to a minimum, creating an environment in which the engrafting stem

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cells will be optimally stimulated. In order for a limiting dilution analysis of the stem cell content of the test cell suspension to be performed, the recipients must be able to survive regardless of whether they receive any stem cells in the test cells injected. Survival of lethally irradiated recipients is ensured by cotransplanting them with hematopoietic cells of the same (host) genotype that contain sufficient numbers of short-term repopulating cells but minimal numbers of long-term repopulating cells. In the case of using c-kit mutant hosts, survival is ensured by pretreating them with a sublethal dose of radiation that allows significant numbers of endogenous cells to survive (Szilvassy et al., 1990; Miller et al., 1996). The differentiated blood cell progeny of the test cells and the recipients must be genetically distinguishable and assessed at a time when they can be safely assumed to represent the exclusive output of cells with life-long stem cell potential. The earliest analysis can be done 1 month post transplantation, but in order to confirm the presence of long-term repopulating cells (or HSCs), donor-derived progeny should be detected at least 4 months post transplantation. Strains of mice congenic with the C57Bl/6 mouse are typically used, to allow the blood cell progeny of the test cells to be uniquely identified by CD45 (Ly5) allotype markers or glucose phosphate isomerase isoform differences. Quantification of HSCs is achieved by application of Poisson statistical analysis on the proportion of animals that test positive for the test cell–derived repopulation at each cell dose transplanted, where the dose at which 37% of animals are negative is estimated to contain 1 HSC or 1 CRU. In practice, a threshold of ≥1% test cell–derived myeloid and lymphoid peripheral blood (PB) cells detected >4 months post-transplant has been shown to rigorously detect a long-term lympho-myeloid repopulating cell. Using this assay, the frequency of HSCs in fresh BM of a mouse has been estimated to be about 1 in 1–2 × 104 nucleated cells (Szilvassy et al., 1990; Rebel et al., 1994), whereas in 5-FU-pretreated BM, the corresponding figure is 1 in 2–5 × 103 (Szilvassy et al., 1989, 1990, 2002). The same strategy and protocols described here could be used to achieve somewhat lower levels of in vitro expansion of murine HSCs by HOXB4 (Antonchuk et al., 2002) or NUP98-HOXB4 (Ohta et al., 2007; i.e., ∼40-fold or ∼300-fold respectively). Moreover, the strategy could be adapted for in vitro HOXB4-mediated HSC expansion together with down-regulation of PBX (Cellot et al., 2007).

Materials 2- to 6-month-old C57Bl/6-W41 /W41 [W41 (Ly5.2)] mice bred and maintained at the British Columbia Cancer Research Centre animal facility according to the Canadian Council on Animal Care (also available from The Jackson Laboratory) Dulbecco’s phosphate-buffered saline without Ca2+ and Mg2+ (CMF-DPBS, StemCell Technologies, cat. no. 37350) containing 2% fetal bovine serum (FBS, StemCell Technologies, cat. no. 06250) Acidified water: prepare 0.1 N HCl in sterile distilled water, then dilute this solution 1:100 in the animals’ drinking water 0.8% (w/v) NH4 Cl/1 mM EDTA in H2 O (StemCell Technologies, cat. no. 07850), ice cold CMF-PBS containing 2% (v/v) fetal bovine serum (FBS, StemCell Technologies, cat. no. 06250); store up to 1 month at 4◦ C Antibodies (fluorochrome-conjugated; BD Pharmingen): B220-PE, Ly6G-PE, Mac1-PE, CD4-PE, CD8-PE, Ly5.1-biotin Streptavidin-APC (BD Pharmingen) CMF-DPBS containing 2% (v/v) FBS and 1 µg/ml propidium iodide Mouse irradiator (X-ray or cesium unit or equivalent) Insulin syringes with 28-G 1/2 -in. needles (BD) Heparinized capillary tubes (e.g., Fisher Scientific) 14-ml polypropylene round-bottom tubes (BD, cat. no. 352059)

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Tabletop centrifuge 96-well U-bottom microtiter plates plate (BD Falcon, cat. no. 353077) 1.4-ml ScreenMates round-bottom storage tubes in a snap rack (Matrix Technologies, cat. no. 4246) L-Calc Software (StemCell Technologies) Additional reagents and equipment for tail-vein injection of the mouse (Donovan and Brown, 2006a), tail-vein blood collection from the mouse (Donovan and Brown, 2006c), and flow cytometry (Robinson et al., 2007) Perform limiting dilution analysis (LDA) of murine CRU 1. Sublethally irradiate W41 (Ly5.2) recipients by exposure to X rays (360 cGy; Miller et al., 1996). 2. Prepare four or five cell mixtures at each cell dose, each in 800 to 1000 µl of icecold CMF-DPBS containing 2% FBS, of each desired dose of test cells and inject 200 µl/recipient intravenously into the lateral tail vain of irradiated W41 recipients (Donovan and Brown, 2006a) using an insulin syringe with a 28-G 1/2 -in. needle. Inject a minimum of three recipients per cell dose. See Table 2A.7.1 as a guide for selecting appropriate test cell doses, expressed as either “starting cell equivalents” (i.e., a constant fraction of the initial culture, regardless of the total nucleated cell output) or as the fraction of total culture.

3. Maintain mice on acidified water for at least 1 month post irradiation.

Assess CRU frequencies in input and cultured murine BM cells 4. Analyze hematopoietic reconstitution of transplanted recipients at any time at least 6 weeks after transplantation. Collect 100 µl of blood from the tail vein of each recipient (as well as nonmanipulated control mouse) into heparinized capillary tubes and flush each blood sample into a 14-ml tube. 5. Lyse erythrocytes by adding 3 ml ice-cold 0.8% NH4 Cl/0.1 mM EDTA), vortex lightly, and incubate on ice 5 to 10 min. Table 2A.7.1 Experimental Details to Assess (by CRU assay) HSC Expansion in Cultures of Murine NUP98-HOXA10HD– Transduced Cells

Initial CRU Numbers of cells assayed at day 10 Postulated content per (starting cell equivalents or fraction day-10 culture (day 0) of individual culture) expansion (fold)

Numbers of cells assayed at day 0

Input per culture (day 0)

3 recipients each to receive 1000, 5000, and 20000 5-FU pretreated BM cells

5-FU BM 3 × 106 cells/10 ml culture

600

3 recipients each to receive equivalent of 2.5, 25, and 250 starting cells

>1000

5-FU BM 5000 cells/100 µl culture

1–2

3 recipients each to receive 1/200 and 1/2000 fractions of individual mini-cultures

>1000

Sca1+ lin− BM 200 cells/100 µl culture c-kit+ Sca1+ lin− BM 30-50 cells/100 µl

1–2 1–2

3 recipients each to receive 1/200th >1000 and 1/2000th fraction of individual mini-cultures

High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

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6. Add 6 ml of CMF-DPBS with 2% FBS, centrifuge 7 min at 300 × g, 4◦ C, and remove the supernatant, leaving no more than 150 to 200 µl. 7. Aliquot ∼50 µl of each blood sample into three separate wells of a 96-well U-bottom microtiter plate. 8. To each triplicate set of blood cells, add 50 µl of antibody dilution (each antibody stock is titrated after purchase: add saturating amounts of antibodies in CMF-DPBS with 2% FBS according to titration) as follows:

biotinylated anti-Ly5.1 in combination with: PE-labeled antibody to B220 (to detect B lymphoid cells) or a combination of PE-labeled antibodies to Ly6G and Mac-1 (to detect myeloid cells) or a combination of PE-labeled antibodies to CD4 and CD8 (to detect T lymphoid cells). Incubate cells for 30 min on ice. Wash all samples with 100 µl per well of CMFDPBS with 2% FBS, centrifuging 7 min at 300 × g, 4◦ C, in a centrifuge fitted with a microtiter plate carrier, and remove supernatants. Incubate an additional 30 min on ice with the appropriate dilution of APC-labeled streptavidin. The second incubation can be avoided by using anti-Ly5.1 directly conjugated to APC. In addition, prepare samples containing unstained cells and cells stained only with PEand APC-conjugated antibodies for establishing threshold and compensation settings on the FACS instrument.

9. Wash all samples after each staining step with 100 µl of CMF-DPBS with 2% FBS and 1 µg/ml propidium iodide, using the centrifugation conditions described in step 8, prior to analysis on a flow cytometric instrument. Transfer samples directly into 1.4-ml plastic round-bottom tubes using a multichannel pipettor. Donor-derived (Ly5.1+ and GFP+ ) myeloid and lymphoid cell populations can be detected using standard flow cytometry procedures (e.g., Ohta et al., 2007).

10. In each group of recipients transplanted with various doses of test cells, determine the proportion of recipients exhibiting at least 1% donor-derived (Ly5.1+ and/or GFP+ ) leukocytes. Score as positive only those recipients in which donor-derived (test) cells are detectable among B (B220+ ) and T (CD4/CD8+ ) lymphoid and myeloid (Ly6G/Mac-1+ ) compartments (see Critical Parameters and Anticipated Results for more details). 11. Determine CRU frequencies by maximum likelihood analysis of the proportions of negative recipients in groups of mice transplanted with various numbers of test cells. Statistical analysis software programs available for this application (L-Calc, StemCell Technologies) are designed to accept three pieces of data: the test cells dose, the total number of mice in each dose group, and number of mice that scored negative at each dose tested.

12. Once CRU frequencies in initial BM population (result of day-0 CRU assay) and at the end of the culture (result of day-10 CRU assay) are measured, estimate CRU content before/after ex vivo expansion, and, therefore, CRU net expansion in the culture (see Anticipated Results for more details).

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

DMEM medium with 15% FBS Dulbecco’s modified Eagle’s medium (high-glucose formulation, 4500 mg glucose/liter (StemCell Technologies, cat. no. 36250) 15% (v/v) fetal bovine serum (FBS; StemCell Technologies cat no. 06250) 1× penicillin/streptomycin (Invitrogen, cat. no. 15140–122) Store up to 1 month at 4◦ C

D-

DMEM wash medium Dulbecco’s modified Eagle’s medium (high-glucose formulation, 4500 mg glucose/liter (StemCell Technologies, cat. no. 36250) 2% (v/v) fetal bovine serum (FBS; StemCell Technologies cat no. 06250) Store up to 1 month at 4◦ C

D-

COMMENTARY Background Information

High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

HSCs appear early in embryogenesis and subsequently amplify their numbers to levels that are then maintained for the lifespan of the individual. During ontogeny, there is a great expansion of all hematopoietic cells, including HSCs, to meet the growing needs of the body. The murine fetal liver at 12 dpc contains approximately 40 HSCs, as detected by the CRU assay. By 16 dpc, this number has expanded 30-fold to 1500 HSCs (Ema et al., 2000), and by adulthood, a further 13-fold expansion brings the total HSC content up to 20,000 (Szilvassy et al., 1990). HSC self-renewal also occurs in the BM following transplantation. There is an initial phase of hematopoietic recovery, when HSCs are stimulated to divide and replenish both primitive and mature hematopoietic compartments. In the murine system, retroviral marking studies, and, more recently, reconstitution studies based on injection of single purified HSC, it has been shown that single HSCs can maintain hematopoiesis for the lifetime of a mouse, and that clones established by HSCs continue to contain newly generated HSC again capable of regenerating the system (Benveniste et al., 2003). The ability to activate HSCs into division without causing their differentiation would be an immensely useful tool both for experimental and clinical uses. One approach that has allowed some HSC expansion in vitro to be achieved has focused on the identification of optimized combinations and concentrations of externally acting growth factors and related molecules (Bryder and Jacobson, 2000;

Bhardwaj et al., 2001; Audet et al., 2002; Varnum-Finney et al., 2003; Willert et al., 2003; Zhang and Lodish, 2005; Nakayama et al., 2006; Zhang et al., 2006a). A complementary approach has been to identify intrinsic regulators such as chromatin modifiers (Ohta et al., 2002; Iwama et al., 2004; Kajiume et al., 2004), key mediators of signaling pathways (Kato et al., 2005; Ema et al., 2005; Zhang et al., 2006b), and transcription factors (Sauvageau et al., 2004; Zeng et al., 2004; Hock et al., 2004; Galan-Caridad et al., 2007) that can be manipulated to activate or promote HSC self-renewal divisions. A striking example of the latter strategy is the use of retrovirally engineered overexpression of the homeobox transcription factor HOXB4 to stimulate expansions of HSC numbers in vitro of up to 80-fold (Antonchuk et al., 2002; Amsellem et al., 2003; Miyake et al., 2006). Moreover, suppression of Pbx1 expression can further enhance in vitro Hoxb4-mediated HSC expansion to a remarkable 100,000-fold (Cellot et al., 2007). Recent studies have suggested the ability of HOXB4 to induce significant expansion of HSCs in culture may extend to other HOX genes. These include results of experiments testing the effect of forced overexpression of HOXA9 (Thorsteinsdottir et al., 1999) and previous data using engineered NUP98-HOX fusion genes (Pineault et al., 2004), showing their ability to block hematopoietic differentiation and to promote the self-renewal of primitive progenitors, as assessed by serial replating of colony-forming cells or expansion of spleen colonies. Remarkable expansions of

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NUP98-HOX–transduced HSCs (300-fold to 10,000-fold over input) in contrast to the expected decline of HSCs in control cultures were discovered by evaluating their presence in 10- to 20-day cultures of transduced mouse BM cells. HSC recovery was measured by limiting-dilution assay for long-term competitive repopulating cells (CRU assay). Importantly, NUP-HOX-expanded HSCs displayed no proliferative senescence and retained normal lympho-myeloid activity and a controlled pool size in vivo. The average magnitude of the HSC expansions achieved by overexpression of the NUP98-HOXB4 fusion gene was ∼300-fold, i.e., ∼4 times the effect of HOXB4 alone using the same vector. The greater than 1000-fold expansions of HSCs obtained using NUP98-HOXA10hd fusion genes are unprecedented and come close to the theoretical limit in a maximum period of 7 to 8 days of gene expression, assuming no significant shortening of the reported 12- to 14-hr cell cycle time for these cells (Habibian et al., 1998; Uchida et al., 2003). Even further levels of HSC expansion could be achieved by extension of the culture period to 17 days, showing that the transduced HSC numbers continued to increase up to a total of greater than 10,000fold (Ohta et al., 2007).

fer, since the right number of producer cells is required at the time of BM harvest. Producer cell numbers should be adjusted according to the plate surface area, in order to achieve 90% confluence (e.g., 6 × 106 cells per 10-cm tissue culture dish). To ensure the optimal infection and further growth of BM cells in culture, it is important to maintain the BM cell concentration at 105 cells/ml (and not more than 106 cells/ml). This is achieved by replating only 10% of the initial culture into the same size dish on day 7. Calculations are done based on one-tenth of the starting number of HSCs. In order to measure HSC frequency in the starting population of 5-FU-pretreated BM cells (day 0) and at the end of the culture period (day 10), CRU assays are to be performed before (day-0 CRU assay) and after expansion (day-10 CRU assay), allowing comparison of before/after CRU contents, and, therefore, estimation of HSC expansion in culture. Recipients of day-0 (nontransduced) or day-10 (NUP98-HOXA10hd-transduced) BM cells whose blood contains greater than 1% donor-derived (Ly5.1+ GFP− or Ly5.1+ GFP+ , respectively) myeloid and lymphoid cells are considered to be positive. All other recipients are scored as negative.

Troubleshooting Critical Parameters Stable integration of murine retroviral vectors requires cell division of the target cells. HSCs, however, are quiescent or cycling very slowly. Therefore, to activate HSCs into cycling, BM donor mice are injected with 5-FU. BM harvested from 5-FU-pretreated mice contains a higher frequency of cycling HSCs susceptible to retroviral infection, and a decreased proportion of more mature cell types. Many groups have established the importance of cytokine stimulation in culture, involving a combination of exposure to growth factors for 24 to 48 hr prior to virus exposure (prestimulation period) and throughout the subsequent period of virus infection (Bodine et al., 1989; Luskey et al., 1992). Cytokines are critical to trigger/maintain cycling and promote survival of HSCs during the infection procedure. Cocultivation of BM cells and retroviral producer cells that have been irradiated usually leads to a higher transduction efficiency than supernatant infection, mainly because the producer cells continuously release viral particles into the culture medium. Planning ahead is very important for this type of gene trans-

See Table 2A.7.2 for troubleshooting information.

Anticipated Results Following 4-day 5-FU treatment, expected cell yield per treated mouse (two femurs and two tibias) is 2–5 × 106 nucleated cells, which is about 10-fold lower than a BM harvest from untreated normal mouse. Cocultivation of 5-FU-pretreated BM cells and irradiated retroviral producer cells consistently leads to more than 75% transduction efficiency by day 10 for all cultures, regardless of the retroviral vector used (MSCV-IRES-GFP or MSCV-NUP98HOXA10hd-IRES-GFP; Ohta et al., 2007). The proliferation rate of control GFP-onlytransduced cells versus NUP98-HOXA10hdtransduced cells is comparable, generating on average a 50- to 100-fold increase in total output per culture by day 10 (i.e., from 3 × 106 initial BM cells to ∼1.5 × 108 , or from 5000 to ∼1 × 106 total nucleated cells by day 10). Nevertheless, in cultures initiated with cells transduced with the control GFP, the total number of GFP+ CRUs present in the 10-day cultures markedly decline (from ∼1 per 5000 starting

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Table 2A.7.2 Troubleshooting Guide for Assessment of Ex Vivo Expansion of Retrovirally Transduced HSCs

Problem

Possible cause

Solution

Lack of 10-fold decrease in number of leukocytes in BM harvested following 4-day 5-FU treatment

1. Presence of residual red blood cell population in the sample. 2. Unsuccessful administration or poor efficacy of the reagent.

1. Repeat lysis and viable cell counts. 2. Ensure the reagent was kept in a dark place and check the expiration date.

Poor transduction efficiency 1. Retroviral producer cells are <90% confluent, thus not producing enough virus. 2. Retroviral producer cells are overconfluent, resulting in decreased virus production. 3. Mycoplasma contamination of retroviral producer cells.

1. Count cells carefully and adjust numbers according to the plate surface area. Use 6 × 106 cells per 10-cm tissue culture dish as a reference. 2. Recalibrate irradiation equipment. 3. Treat for mycoplasma infection.

Disruption of adherent 1. Use of cold medium for washing. retroviral producer cells 2. Vigorous washing. during the removal of BM at the end of cocultivation

1. Use prewarmed (37◦ C) medium to wash BM from retroviral producer cells. 2. Wash as gently as possible.

Poor CRU recovery in day-10 expanded cultures

HSCs are often strongly attached to the producer cells

Wash BM from producer cells at least 5 times to get as many BM as possible

Problems with mouse injections/bleeds

Tail veins are not dilated

Gently warm mice for 3-5 min using a heat lamp and thoroughly clean their tails with alcohol pads to make veins more visible

cells to less than 1 per 200,000 starting cell equivalents on day 10, i.e., an absolute decrease of ∼50-fold). In contrast, the number of NUP98-HOXA10hd-transduced CRUs significantly increases during the same period (to ∼1 per 5 starting cell equivalents, an expansion of ∼1000-fold). Given the huge decline in survival of cultured HSCs if not transduced with NUP98-HOXA10hd, the complete concordance between the presence of regenerated donor-derived cells (Ly5.1+ ) and transduced (GFP+ ) cells in the reconstituted recipients is expected.

Time Considerations The time required for generation of NUP98-HOX–expanded HSCs in culture includes 4 days of 5-FU treatment, 2 days of prestimulation, 2 days of infection, and at least 6 days of expansion period (at least 14 days total). Long-term lympho-myeloid reconstitution of recipients transplanted with NUP98-HOXexpanded HSCs should be analyzed 4 months after transplantation. High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

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E., Sasaki, Y., and Jacobsen, S.E.W. 2001. Upregulation of flt3 expression within the bone marrow Lin− Sca1+ c-kit+ stem cell compartment is accompanied by loss of self-renewal capacity. Immunity 15:659-669. Amsellem, S., Pflumio, F., Bardinet, D., Izac, B., Charneau, P., Romeo, P.H., DubartKupperschmitt, A., and Fichelson, S. 2003. Ex vivo expansion of human hematopoietic stem cells by direct delivery of the HOXB4 homeoprotein. Nat. Med 9:1423-1427. Antonchuk, J., Sauvageau, G., and Humphries, R.K. 2002. HOXB4-induced expansion of adult hematopoietic stem cells ex vivo. Cell 109:3945. Audet, J., Miller, C.L., Eaves, C.J., and Piret, J.M. 2002. Common and distinct features of cytokine effects on hematopoietic stem and progenitor cells revealed by dose-response surface analysis. Biotechnol. Bioeng. 80:393-404. Balazs, A.B., Fabian, A.J., Esmon, C.T., and Mulligan, R.C. 2006. Endothelial protein C receptor (CD201) explicitly identifies hematopoietic stem cells in murine bone marrow. Blood 107:2317-2321. Benveniste, P., Cantin, C., Hyam, D., and Iscove, N.N. 2003. Hematopoietic stem cells engraft in mice with absolute efficiency. Nat. Immunol. 4:708-713. Bhardwaj, G., Murdoch, B., Wu, D., Baker, D.P., Williams, K.P., Chadwick, K., Ling, L.E., Karanu, F.N., and Bhatia, M. 2001. Sonic hedgehog induces the proliferation of primitive human

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hematopoietic cells via BMP regulation. Nat. Immunol. 2:172-180. Bodine, D.M., Karlsson, S., and Nienhuis, A.W. 1989. Combination of interleukins 3 and 6 preserves stem cell function in culture and enhances retrovirus-mediated gene transfer into hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 86:8897-8901. Bodine, D.M., McDonagh, K.T., Seidel, N.E., and Nienhuis, A.W. 1991. Survival and retrovirus infection of murine hematopoietic stem sell in vitro: Effects of 5-FU and method of infection. Exp. Hematol. 19:206-212. Bryder, D. and Jacobsen, S.E.W. 2000. Interleukin3 supports expansion of long-term multilineage repopulating activity after multiple stem cell divisions in vitro. Blood 96:1748-1755. Cellot, S., Krosl, J., Chagraoui, J., Meloche, S., Humphries, R.K., and Sauvageau, G. 2007. Sustained in vitro trigger of self-renewal divisions in Hoxb4hiPbx1(lo) hematopoietic stem cells. Exp. Hematol. 35:802-816. Chen, C.Z., Li, M., de Graaf, D., Monti, S., Gottgens, B., Sanchez, M.J., Lander, E.S., Golub, T.R., Green, A.R., and Lodish, H.F. 2002. Identification of endoglin as a functional marker that defines long-term repopulating hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 99:15468-15473. Christensen, J.L. and Weissman, I.L. 2001. Flk-2 is a marker in hematopoietic stem cell differentiation: A simple method to isolate long-term stem cells. Proc. Natl. Acad. Sci. U.S.A. 98:1454114546. Donovan, J.D. and Brown, P. 2006a. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Donovan, J.D. and Brown, P. 2006b. Parenteral injections. Curr Protoc. Immunol. 73:1.6.11.6.10. Donovan, J.D. and Brown, P. 2006c. Blood collection. Curr. Protoc. Immunol. 73:1.7.1-1.7.9. Ema, H., Takano, H., Sudo, K., and Nakauchi, H. 2000. In vitro self-renewal division of hematopoietic stem cells. J. Exp. Med. 192:1281-1288. Ema, H., Sudo, K., Seita, J., Matsubara, A., Morita, Y., Osawa, M., Takatsu, K., Takaki, S., and Nakauchi, H. 2005. Quantification of selfrenewal capacity in single hematopoietic stem cells from normal and Lnk-deficient mice. Dev. Cell 8:907-914. Galan-Caridad, J.M., Harel, S., Arenzana, T.L., Hou, Z.E., Doetsch, F.K., Mirny, L.A., and Reizis, B. 2007. Zfx controls the self-renewal of embryonic and hematopoietic stem cells. Cell 129:345-357. Habibian, H.K., Peters, S.O., Hsieh, C.C., Wu, J., Vergilis, K., Grimaldi, C.I., Carlson, J.E., Frimberger, A.E., Stewart, F.M., and Quesenberry, P.J. 1998. The fluctuating phenotype of the lymphohematopoietic stem cell with cell cycle transit. J. Exp. Med. 188:393-398.

Harrison, D.E. and Lerner, C.P. 1991. Most primitive hematopoietic stem cells are stimulated to cycle rapidly after treatment with 5-fluorouracil. Blood 78:1237-1240. Hock, H., Meade, E., Medeiros, S., Schindler, J.W., Valk, P.J., Fujiwara, Y., and Orkin, S.H. 2004. Tel/Etv6 is an essential and selective regulator of adult hematopoietic stem cell survival. Genes Dev. 18:2336-2341. Iwama, A., Oguro, H., Negishi, M., Kato, Y., Morita, Y., Tsukui, H., Ema, H., Kamijo, T., Katoh-Fului, Y., Koseki, Y., van Lohuizen, M., and Nakauchi, H. 2004. Enhanced selfrenewal of hematopoietic stem cells mediated by the polycomb gene product Bmi-1. Immunity 21:843-851. Kajiume, T., Ninomiya, Y., Ishihara, H., Kanno, R., and Kanno, M. 2004. Polycomb group gene mel18 modulates the self-renewal activity and cell cycle status of hematopoietic stem cells. Exp. Hematol. 32:571-578. Kalberer, C., Antonchuk, J., and Humphries, R.K. 2002. Genetic modification of murine hematopoietic stem cells by retroviruses In Hematopoietic Stem Cell Protocols (C.A. Klug and C.T. Jordan, eds.) pp. 231-242. Humana Press, Totowa, N.J. Kasper, L.H., Brindle, P.K., Schnabel, C.A., Pritchard, C.E., Cleary, M.L., and van Deursen, J.M. 1999. CREB binding protein interacts with nucleoporin-specific FG repeats that activate transcription and mediate NUP98-HOXA9 oncogenicity. Mol. Cell Biol. 19:764-776. Kato, Y., Iwama, A., Tadokoro, Y., Shimoda, K., Minoguchi, M., Akira, S., Tanaka, M., Miyajima, A., Kitamura, T., and Nakauchi, H. 2005. Selective activation of STAT5 unveils its role in stem cell self-renewal in normal and leukemic hematopoiesis. J. Exp. Med. 202:169179. Kiel, M.J., Yilmaz, O.H., Iwashita, T., Yilmaz, O.H., Terhorst, C., and Morrison, S.J. 2005. SLAM family receptors distinguish hematopoietic stem and progenitor cells and reveal endothelial niches for stem cells. Cell 121:11091121. Luskey, B.D., Rosenblatt, M., Zsebo, K., and Williams, D.A. 1992. Stem cell factor, interleukin-3, and interleukin-6 promote retroviral-mediated gene transfer into murine hematopoietic stem cells. Blood 80:396-402. Matsuzaki, Y., Kinjo, K., Mulligan, R.C., and Okano, H. 2004. Unexpectedly efficient homing capacity of purified murine hematopoietic stem cells. Immunity 20:87-93. Miller, C.L., Rebel, V.I., Lemieux, M.E., Helgason, C.D., Lansdorp, P.M., and Eaves, C.J. 1996. Studies of W mutant mice provide evidence for alternate mechanisms capable of activating hematopoietic stem cells. Exp. Hematol. 24:185194. Miyake, N., Brun, A.C., Magnusson, M., Miyake, K., Scadden, D.T., and Karlsson, S. 2006.

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HOXB4-induced self-renewal of hematopoietic stem cells is significantly enhanced by p21 deficiency. Stem Cells 24:653-661. Nakayama, A., Matsui, H., Fukushima, T., Ichikawa, H., Yamada, K., Amao, T., Hosono, M., and Sugimoto, K. 2006. Murine serum obtained from bone marrow–transplanted mice promotes the proliferation of hematopoietic stem cells by co-culture with MS-5 murine stromal cells. Growth Factors 24:55-65. Ohta, H., Sawada, A., Kim, J.Y., Tokimasa, S., Nishiguchi, S., Humphries, R.K., Hara, J., and Takihara, Y. 2002. Polycomb group gene rae28 is required for sustaining activity of hematopoietic stem cells. J. Exp. Med. 195:759-770. Ohta, H., Sekulovic, S., Bakovic, S., Eaves, C.J., Pineault, N., Gasparetto, M., Smith, C., Sauvageau, G., and Humphries, R.K. 2007. Near-maximal expansion of hematopoietic stem cells in culture using NUP98-HOX fusions. Exp. Hematol. 35:817-830. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Pineault, N., Abramovich, C., Ohta, H., and Humphries, R.K. 2004. Differential and common leukemogenic potentials of multiple NUP98-Hox fusion proteins alone or with Meis 1. Mol. Cell Biol. 24:1907-1917. Rebel, V.I., Dragowska, W., Eaves, C.J., Humphries, R.K., and Lansdorp, P.M. 1994. Amplification of Sca-1+ Lin− WGA+ cells in serum-free cultures containing steel factor, interleukin-6, and erythropoietin with maintenance of cells with long-term in vivo reconstituting potential. Blood 83:128-136. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. (eds.) 2007. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Sauvageau, G., Iscove, N.N., and Humphries, R.K. 2004. In vitro and in vivo expansion of hematopoietic stem cells. Oncogene 23:72237232. Shizuru, J.A., Negrin, R.S., and Weissman, I.L. 2005. Hematopoietic stem and progenitor cells: Clinical and preclinical regeneration of the hematolymphoid system. Annu. Rev. Med. 56:509-538. Szilvassy, S.J., Lansdorp, P.M., Humphries, R.K., Eaves, A.C., and Eaves, C.J. 1989. Isolation in a single step of a highly enriched murine hematopoietic stem cell population with competitive long-term repopulating ability. Blood 74:930-939.

High Level In Vitro Expansion of Murine Hematopoietic Stem Cells

Szilvassy, S.J., Humphries, R.K., Lansdorp, P.M., Eaves, A.C., and Eaves, C.J. 1990. Quantitative assay for totipotent reconstituting hematopoietic stem cells by a competitive repopulation strategy. Proc. Natl. Acad. Sci. U.S.A. 87:87368740.

Szilvassy, S., Nicolini, F., Eaves, C., and Miller, C. 2002. Quantitation of murine and human hematopoietic stem cells by limiting-dilution analysis in competitive repopulated hosts. In Hematopoietic Stem Cell Protocols (C.A. Klug and C.T. Jordan, eds.) pp. 167-187. Humana Press, Totowa, N.J. Thorsteinsdottir, U., Sauvageau, G., and Humphries, R.K. 1999. Enhanced in vivo regenerative potential of HOXB4-transduced hematopoietic stem cells with regulation of their pool size. Blood 94:2605-2612. Uchida, N., Dykstra, B., Lyons, K.J., Leung, F.Y.K., and Eaves, C.J. 2003. Different in vivo repopulating activities of purified hematopoietic stem cells before and after being stimulated to divide in vitro with the same kinetics. Exp. Hematol. 31:1338-1347. Varnum-Finney, B., Brashem-Stein, C., and Bernstein, I.D. 2003. Combined effects of Notch signaling and cytokines induce a multiple log increase in precursors with lymphoid and myeloid reconstituting ability. Blood 101:17841789. Verma, I.M. and Weitzman, M.D. 2005. Gene therapy: Twenty-first century medicine. Annu. Rev. Biochem. 74:711-738. Wagers, A.J. and Weissman, I.L. 2006. Differential expression of alpha2 integrin separates long-term and short-term reconstituting Lin-/ loThy1.1(lo)c-kit+ Sca-1+ hematopoietic stem cells. Stem Cells 24:1087-1094. Willert, K., Brown, J.D., Danenberg, E., Duncan, A.W., Weissman, I.L., Reya, T., Yates, J.R. III, and Nusse, R. 2003. Wnt proteins are lipidmodified and can act as stem cell growth factors. Nature 42:448-452. Zeng, H., Yucel, R., Kosan, C., Klein-Hitpass, L., and Moroy, T. 2004. Transcription factor Gfi1 regulates self-renewal and engraftment of hematopoietic stem cells. EMBO J. 23:41164125. Zhang, C.C. and Lodish, H.F. 2005. Murine hematopoietic stem cells change their surface phenotype during ex vivo expansion. Blood 105:4314-4320. Zhang, C.C., Kaba, M., Ge, G., Xie, K., Tong, W., Hug, C., and Lodish, H.F. 2006a. Angiopoietinlike proteins stimulate ex vivo expansion of hematopoietic stem cells. Nat. Med. 12:240245. Zhang, J., Grindley, J.C., Yin, T., Jayasinghe, S., He, X.C., Ross, J.T., Haug, J.S., Rupp, D., PorterWestpfahl, K.S., Wiedemann, L.M., and Wu, H., Li, L. 2006b. PTEN maintains haematopoietic stem cells and acts in lineage choice and leukaemia prevention. Nature 441:518522.

2A.7.14 Supplement 4

Current Protocols in Stem Cell Biology

Isolation and Visualization of Mouse Placental Hematopoietic Stem Cells

UNIT 2A.8

Christos Gekas,1 Katrin E. Rhodes,1 and Hanna K.A. Mikkola1 1

University of California Los Angeles, Los Angeles, California

ABSTRACT This unit describes the isolation of hematopoietic stem cells (HSCs) from the mouse placenta. The placenta was recently identified as an important hematopoietic site that generates HSCs de novo and provides a transitory niche for a large pool of HSCs during midgestation. This protocol includes a dissection technique for murine placenta, the mechanical and enzymatic steps of placental tissue dissociation, and phenotypical identification and isolation of HSCs. It also contains a method for immunohistochemical analysis of placenta tissue sections to visualize developing HSCs in the placenta. Curr. C 2008 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 6:2A.8.1-2A.8.14.
Keywords: hematopoietic stem cell (HSC) r placenta r fetal r dissection r flow cytometry r FACS sorting r fixed-frozen sections r immunohistochemistry

INTRODUCTION This unit describes the isolation of hematopoietic stem cells (HSCs) from the placenta and a method for visualizing nascent HSCs and hematopoietic progenitors by immunohistochemistry. The authors and others recently identified the placenta as an important fetal hematopoietic site that generates HSCs de novo and provides a transitory niche for a large pool of HSCs during midgestation (Gekas et al., 2005; Ottersbach and Dzierzak, 2005; Rhodes et al., 2008). The unit begins with a detailed description of techniques for the dissection of murine placenta (Basic Protocol 1), followed by protocols for dissociation of placental tissue into single-cell suspensions (Basic Protocol 2), isolation of placental HSCs by FACS sorting (Basic Protocol 3), and visualization of placental HSCs by immunohistochemistry (Basic Protocol 4 and Support Protocol).

DISSECTION OF MURINE PLACENTA This protocol is used for dissection of placentas from murine embryos. All dissections are performed under an inverted dissection microscope. Depending on the age of the embryo, slightly different dissection techniques are used, as outlined below.

BASIC PROTOCOL 1

NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. NOTE: If placental cells are to be used for culturing or transplantation, all solutions and equipment should be kept sterile, and proper aseptic technique should be used accordingly.

Materials Pregnant mouse with midgestation embryos Isoflurane 1× Dulbecco’s phosphate-buffered saline with calcium and magnesium (DPBS; Cellgro/Mediatech) Current Protocols in Stem Cell Biology 2A.8.1-2A.8.14 Published online August 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a08s6 C 2008 John Wiley & Sons, Inc. Copyright


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Figure 2A.8.1 Collecting embryos from a pregnant dam. (A) The uterine horns are picked up with forceps and cut out with scissors. (B) The conceptuses are placed in a petri dish and the endometrium is peeled away with forceps.

1× DPBS (with calcium and magnesium; Cellgro/Mediatech)/5% (v/v) fetal bovine serum (FBS) Scissors Number 5 or 55 dissection forceps, stainless steel or titanium 35-mm petri dishes Ice Inverted dissection microscope 15-ml centrifuge tubes 1. Sacrifice the mice by isoflurane inhalation. Collect the embryos from the timed pregnant dam by making a small incision in the lower abdominal area and tearing away the skin with both hands to uncover the abdomen. 2. Cut open the peritoneum. Using forceps, pick up the uterine horns and remove them using scissors at each distal end (Fig. 2A.8.1A). Place the uterine horns in a petri dish on ice and wash several times with DPBS. 3. Using an inverted dissection microscope and two forceps, begin to carefully peel away the endometrial tissue surrounding the conceptus (see Fig. 2A.8.1B; see Video 1). Care should be taken not to puncture or otherwise inflict structural damage on the conceptuses, as this will complicate subsequent dissection steps.

4. Place the isolated conceptuses in a new petri dish with DPBS on ice. 5. Transfer one conceptus to a new petri dish with DPBS placed under the microscope and peel away the decidua from the placenta using two forceps (E10.5: Fig. 2A.8.2A to D; E12.5: Fig. 2A.8.2E to H; also see Fig. 2A.8.5; see Videos 2 and 3). NOTE: Removal of decidua is optional before immunohistochemistry. The thickness of the decidua layer relative to the size of the placenta decreases with age. Until E10.5, the decidua surrounds the entire conceptus (E10.5; Fig. 2A.8.2A), whereas at later developmental stages it becomes progressively thinner and covers only the maternal side of the placenta (E12.5; Fig. 2A.8.2E).

Placental Hematopoietic Stem Cells

Care should be taken to remove as much as possible of the decidua from the placenta in order to minimize aggregation of cells during dissociation into a single-cell suspension. Furthermore, decidual cells are both highly autofluorescent and express some of the known HSC markers; therefore, removal of decidual cells before flow cytometry allows for more accurate identification of HSCs. A smooth continuous peeling motion is recommended.

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Figure 2A.8.2 Removing the decidua of E10.5 (A to D) and E12.5 (E to H) embryos. (A) At E10.5 the decidua surrounds the entire conceptus. (B to C) The decidua (white) is peeled away with forceps. (D) The placenta and the giant cell layer surrounding the embryo (top) after successful removal of the decidua (bottom). (E) At E12.5 the decidua is thinner and covers the endometrial side of the placenta. (F to G) The decidua (white) is peeled away with forceps. (H) The embryo (right) after successful removal of the decidua (left). Magnification: 15×.

Figure 2A.8.3 Removing the yolk sac and detaching the placenta from an E12.5 embryo. (A) The yolk sac has been cut away from the chorionic plate (arrow) and moved to the other side of the embryo. (B,C) Using forceps, the umbilical cord (arrow) is detached from the placenta. (D) The placenta (top left) is dissected out from the embryo (top right) and the yolk sac (bottom). Magnification: (A, D) 15×; (B,C) 22×.

6. Cut the yolk sac from the placenta at the junction between the two organs and gently pull the yolk sac and vitelline vessels away from the placenta (E10.5 and E12.5, respectively; see Videos 4 and 5). 7. Hold the chorionic plate carefully with one of the forceps; hold the umbilical cord as close as possible to the chorionic plate with the other forceps. Pull the umbilical cord and attached embryo away from the placenta (E12.5: Fig. 2A.8.3A to D; see Video 6). Care should be taken not to disrupt the chorionic plate, especially at earlier developmental stages (E8.5 to 10.5). Disrupting the chorionic plate can result in loss of HSCs or damage to the morphology of the placenta.

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8. Cut the excess giant cell tissue at the edges of the placenta. See Video 7. This step is recommended in order to minimize cell clumping during preparation of single-cell suspensions.

9. Collect the placentas in a suitable container, such as a 15-ml conical tube, with ∼500 ml DPBS/5% FBS on ice. One placenta or all of the placentas from one litter may be placed in a tube, depending on the expected genotypes and the experimental question. Depending on the purpose of the experiment, placentas can be dissociated to acquire a single-cell suspension (Basic Protocol 2) for flow cytometry and isolation of HSCs by FACS sorting (Basic Protocol 3), or they can be used for sectioning and immunohistochemical analysis (Basic Protocol 4 and Support Protocol). BASIC PROTOCOL 2

DISSOCIATION OF PLACENTAL TISSUE The following steps are performed to obtain a single-cell suspension of placental tissue to use for subsequent analyses by flow cytometry and functional hematopoietic assays (e.g., colony forming assay) and for transplantation. The process involves the dissociation of the tissue through a series of mechanical and enzymatic steps. NOTE: If placental cells are to be used for culturing or transplantation, all solutions and equipment should be kept sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Materials 1% (w/v) collagenase stock solution: prepared by dissolving collagenase type IA (from Clostridium histolyticum) lyophilized powder (Sigma) in water; dispense into aliquots and store at −20◦ C 1× Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS; Cellgro/Mediatech) Fetal bovine serum (FBS) 10,000 U penicillin and 10,000 µg streptomycin/ml 0.85% saline (P/S; Invitrogen) Dissected placentas (Basic Protocol 1) 15-ml conical tubes 5-ml plastic syringes (BD) 16-G, 18-G, 20-G, 22-G, and 25-G needles (BD) 50-µm cell filters (CellTrics, Partec) 1. Prepare a 0.1% (w/v) collagenase from 1% stock solution by diluting in CMF-DPBS with 10% (v/v) FBS and 1% (v/v) P/S. Fetal calf serum (FCS) can be used as well.

2. Centrifuge the placentas from Basic Protocol 1, step 9, 5 min at 300 × g, 4◦ C, and aspirate the supernatant. 3. Add the collagenase solution to the placentas. The amount of collagenase solution needed varies with embryonic age and number of placentas. A volume that is at least twice the volume of the placentas and between 1 and 5 ml is recommended. Placental Hematopoietic Stem Cells

4. Use the 5-ml syringe and 16-G needle to mechanically disrupt the tissue by passing the collagenase solution and placenta through the needle three times. Repeat with an 18-G needle.

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5. Incubate the sample 45 min at 37◦ C. 6. Pass the sample through a 20-G needle, three times, and incubate for an additional 45 min. Performing this passaging after the initial 45-min incubation has been found to increase HSC yield.

7. Pass the cell solution three times each through 22-G and 25-G needles, sequentially. 8. Filter the sample through a 50-µm filter into a new 15-ml conical tube. Centrifuge 5 min at 300 × g, 4◦ C. Note that cell clumping can occur at this point. These clumps can be difficult to dissociate, as they tend to re-aggregate even after filtering and should therefore be removed from the sample.

9. Aspirate and discard the supernatant. Wash two times by adding 5 ml CMFDPBS/5%FBS and centrifuging 5 min at 300 × g, 4◦ C, aspirating and discarding the supernatant each time.

ISOLATION OF PLACENTAL HSCs BY FACS SORTING The following protocol describes a method for isolation of hematopoietic stem cells and progenitors from placenta by staining with fluorescent-labeled antibodies and FACS sorting. Antibodies and solutions should be kept on ice.

BASIC PROTOCOL 3

Materials Dissociated placental cells (Basic Protocol 2) 1× Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS; Cellgro/Mediatech) 1× red blood cell (RBC) lysis buffer (eBiosciences) Phycoerythrin-labeled rat-anti-mouse c-kit antibody (c-kit-PE; BD Pharmingen) Allophycocyanin-labeled rat-anti-mouse CD34 antibody (CD34-APC; eBiosciences) 7-amino-actinomycin D (7-AAD; BD Biosciences) Tissue-culture treated, round-bottom 96-well plate (BD; Falcon) or 15-ml conical tube or 1.5-ml microcentrifuge tube Centrifuge with rotor adapted for 96-well plates, optional FACS tubes (5-ml 12 × 75–mm polypropylene round-bottom test tube, e.g., BD Falcon) Flow cytometer or a cell sorter (e.g., LSR II, BD; FACSAria, BD) 1. Resuspend the dissociated placental cells from Basic Protocol 2, step 9, in an appropriate volume (depending on number of stains used) of CMF-DPBS and transfer to a 96-well plate or to another suitable container (e.g., a 15-ml conical tube or microcentrifuge tube). Depending on the number of stains and required controls to be performed, use of a 96well plate can facilitate staining, and the resuspension volume will have to be modified accordingly. The maximum recommended volume for the 96-well plate wells is 200 µl.

2. Centrifuge the cells 5 min at 300 × g, 4◦ C, and aspirate or discard the supernatant. All wells of a 96-well plate can be decanted at once by removing the lid and quickly turning the plate upside down. To avoid losing cells, this movement should only be done once.

3. Perform red cell lysis using the 1× RBC lysis buffer per manufacturer’s instructions. In brief, this involves adding ice-cold RBC lysis buffer to the cell pellet, resuspending it by pipetting, and incubating 5 min on ice.

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If performing this step on a 96-well plate, use 50 µl of 1× RBC lysis buffer. In microcentrifuge tubes or 15-ml conical tubes, up to 1000 µl can be used. Red cell lysis is recommended when performing flow cytometry on placental cells, due to the large amount of enucleated material from red cells that would otherwise interfere with the analysis. It is not required if the cells are used directly for colony assays or transplantation. Red cell lysis buffer contains ammonium chloride, pH 7.2, and it can also be prepared in the lab instead of purchasing it.

4. Add an appropriate volume of CMF-DPBS (e.g., up to 200 µl for 96-well plates or ∼10 ml for a 15-ml centrifuge tube) to terminate the reaction. Centrifuge the cells 5 min at 300 × g, 4◦ C. Aspirate and discard the supernatant and wash with another aliquot of CMF-DPBS. 5. Prepare an antibody staining solution by mixing c-kit-PE and CD34-APC in CMFDPBS. For <1 to 2 million cells, a staining volume of 50 µl is recommended; otherwise scale up accordingly. The antibody dilutions should be determined through titration; however for this stain we used a dilution of 1:200 for c-kit-PE and 1:50 for CD34-APC. Although beyond the scope of this protocol, proper flow cytometry controls for autofluorescence, unspecific binding, and compensation should be carried out.

6. Add the antibody dilution to the cells, and mix well by pipetting. Incubate 20 min on ice in the dark. 7. Centrifuge 5 min at 300 × g, 4◦ C. 8. Aspirate and discard the supernatant. Wash three times by adding an appropriate volume of CMF-DPBS (e.g., up to 200 µl for 96-well plates or 15 ml for a 15-ml centrifuge tube) and centrifuging 5 min at 300 × g, 4◦ C, aspirating and discarding the supernatant each time. 9. Prepare a 1:50 dilution of 7-AAD in 200 µl CMF-DPBS (per sample) and resuspend the cells in this solution after the final wash. 7-AAD is used as a viability marker, as it intercalates into the DNA of dying or dead cells but is excluded by viable cells. It is important to exclude dead cells in flow cytometry because their autofluorescence could otherwise be interpreted as false-positive signals. 7-AAD signal is detected on the same channel as PerCP-Cy5.5. Other viability markers (e.g., TO-PRO-1, same channel as FITC) could be used alternatively.

Figure 2A.8.4 Identification of placental HSCs/progenitors by flow cytometry. FACS plots of placenta gating on forward and side scatter (FSC-A/SSC-A) for hematopoietic cells (left), exclusion of 7-AAD-expressing dead cells (middle) and expression of CD34-APC and c-kit-PE (right). HSCs and multipotential progenitor cells are double positive for CD34 and c-kit (outlined in blue).

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10. Transfer the samples to 5-ml FACS tubes and run them in a flow cytometer or a cell sorter. The HSCs and multipotential progenitors are found in the 7-AAD-negative, c-kit-PE-high, and CD34-APC-dim fraction (Fig. 2A.8.4).

PREPARATION OF PLACENTAL TISSUE FOR IMMUNOHISTOCHEMISTRY

BASIC PROTOCOL 4

The following protocol is designed to ensure proper fixation and cryopreservation of placental tissue before embedding the tissue in frozen blocks for immunohistochemistry. This protocol also describes how to orient the placenta properly when preparing the blocks.

Materials Dissected placentas (Basic Protocol 1) 1× Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS; Cellgro/Mediatech), 4◦ C 4% (w/v) paraformaldehyde (PFA): dispense into aliquots and store at −20◦ C 30% (w/v) sucrose solution Optimal cutting temperature solution (O.C.T., Tissue-Tek) Dry ice Drierite Forceps Tissue culture–treated, round-bottom 96-well or 24-well plates (BD Falcon) Razor blade 10 × 10 × 5–mm disposable vinyl specimen molds (e.g., Cryomolds, Tissue-Tek) Permanent marker Small plastic bags −80◦ C freezer Fix placental tissue 1. After isolating whole placenta tissue as described in Basic Protocol 1, step 9, place the tissue in cold (4◦ C) CMF-DPBS in either a 96- or 24-well plate or a 35-mm petri dish. 2. Using forceps, transfer each placenta into a well of a 96- or 24-well plate filled with freshly thawed 4% PFA. Fix for 2 to 4 hr at 4◦ C, depending on the age and size of the tissue. NOTE: It is important that the placenta is fully immersed. If the placentas are very small, you can remove the PBS carefully with a pipet and add the PFA into the same well.

Cryopreserve placental tissue 3. Transfer the tissue into another well containing 30% sucrose solution. Let the tissues incubate overnight at 4◦ C. The tissue must be fully immersed and should ultimately sink to the bottom of the well, indicative of proper cryopreservation. If the placentas are very small you can remove the PFA carefully with a pipet and add the sucrose into the same well.

4. Remove half of the sucrose solution, replace the volume with O.C.T., and place the tissue back at 4◦ C for 1 to 2 hr. This facilitates O.C.T. penetration.

5. Transfer the tissue to 100% O.C.T. and incubate 1 hr at room temperature.

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Embed placental tissue 6. Label the plastic molds with appropriate tissue identification information, using a permanent marker. 7. Pull the tissue out of the O.C.T. solution and cut the placenta in half: orient the disc-shaped placenta with the umbilical cord side facing down and carefully slice in half with a clean razor blade. Move the blade back and forth before releasing pressure to achieve a clean cut. 8. Place each half of the placenta side by side at the bottom of the mold with the cut edge in contact with the bottom surface of the mold. 9. Slowly pour/drip O.C.T. solution into the mold until it is completely full. Avoid producing any bubbles. It can be difficult to keep the placentas in the proper orientation, hold one tissue half in place with forceps and rest the other tissue half against the outside of the forceps, then add the O.C.T.

10. Immediately, place the mold onto dry ice to flash freeze the tissue. The O.C.T. will turn white. After placement of the first block onto dry ice, a small indentation is imprinted into the ice. Use this indentation to stabilize the molds when flash freezing additional placentas.

11. Place the mold into a small plastic bag containing Drierite and store in a −80◦ C freezer. 12. Prepare 5- to 8-µm sections of the placental tissue using a cryotome. Mount the sections on a glass microscope slide. SUPPORT PROTOCOL

IMMUNOHISTOCHEMISTRY ON FIXED FROZEN PLACENTAL TISSUE SECTIONS FOR HSCs This protocol is designed to stain placental tissue sections for a marker of nascent HSCs and progenitors (CD41) and landmarks of the placental structures, endothelium (CD31) and trophoblast (cytokeratin; Fig. 2A.8.5). The fixed frozen blocks must be sectioned into 5-µm to 8-µm thick sections, and each section should contain both the chorionic plate and labyrinth (see Fig. 2A.8.5A). NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Placental Hematopoietic Stem Cells

Materials 5- to 8-µm placental sections mounted on glass microscope slides (Basic Protocol 4) 10× GO buffer (see recipe) 1000 U/ml glucose oxidase (GO; Sigma) 1× Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS; Cellgro/Mediatech) 2.5 µg/ml proteinase K solution: dilute 20 mg/ml proteinase K solution (Amresco) in CMF-DPBS (Cellgro/Mediatech); store up to 3 months at 4◦ C Tyramide amplification kit (Invitrogen) including: Tyramide blocking buffer Streptavidin (SA)-horseradish peroxidase (HRP) CD41 primary antibody (BD Pharmingen) Biotinylated anti-rat IgG (Vector Laboratories) secondary antibody CMF-DPBS/0.1% (v/v) Tween 20

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Vectastain ABC-AP kit (alkaline phosphatase–based conjugate) or ABC Elite kit (peroxidase-based conjugate): each kit includes reagent A and reagent B (Vector Laboratories) Appropriate substrate solutions: peroxidase substrate kit DAB (Vector Laboratories) or alkaline phosphatase substrate kit III (Vector Blue, Vector Laboratories) or alkaline phosphatase substrate kit I (Vector Red, Vector Laboratories) Levamisole (Vector Laboratories) CD31/Pecam primary antibody (BD Pharmingen) Cytokeratin primary antibody (Dako Cytomation) Normal blocking buffer: 5% (v/v) normal horse serum (Vector Laboratories)/ 0.05% (v/v) Tween 20 (Ultra, Sigma)/PBS Avidin/biotin blocking kit (Vector Laboratories) Biotinylated anti-rabbit IgG (Vector Laboratories) Mounting medium, aqueous-based mounting medium (e.g., Vectamount AQ, Vector Laboratories) Plastic microscope slide mailers (Fisher) 2.5-mm Super Pap Pen HT slide markers (Research Products International, Fisher) Humidified chamber (plastic slide box with 1/4 in. of water), recommended Vacuum apparatus: glass filter flask with vacuum side arms and plastic tubing (Nalgene 180 PVC, nontoxic and autoclavable) Transfer pipets Microscope Prepare sections for immunohistochemistry 1. Incubate slides with placental sections 30 min at 37◦ C. 2. Dilute 10× GO buffer to 1× with CMF-DPBS and warm by incubating 30 min at 37◦ C. CAUTION: GO buffer is toxic! Handle with care.

3. After 30 min dilute the 1000 U/ml glucose oxidase 1:1000 in the prewarmed buffer and place in a slide mailer. Approximately 30 ml is required per plastic slide mailer (which holds five slides). Either a slide mailer or a Coplin jar may be used for these steps, but the volume of reagents required for the slide mailer is smaller than for a Coplin jar. In addition, the slide mailer closes tightly, an advantage in incubation steps.

4. Wash the slides three times in CMF-DPBS by transferring them to slide mailers with fresh solution. 5. Incubate the slides in the diluted glucose oxidase for 30 min at 37◦ C. Incubating the placental sections with glucose oxidase quenches endogenous peroxidase activity that would interfere with staining.

6. Wash the slides three times in CMF-DPBS. 7. Use a Pap-Pen to encircle each tissue section on the slide. This creates a wax border around the tissue. The wax should not touch the tissue. The following steps in this protocol are performed using a small volume of reagent for each tissue spot. The reagent should be enough to cover the tissue section forming a small bubble within the wax circle. If the tissue and wax circle are difficult to see, it is helpful to draw a circle (with a permanent marker) on the bottom side of the slide around the tissue.

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Retrieve antigen 8. Carefully add ∼100 µl of 2.5 µg/ml proteinase-K solution to each tissue spot for 7 to 10 min at room temperature. If the tissue spots are large, a greater volume of proteinase-K can be used. The tissue must be fully covered. For this and the remaining steps it is recommended that the slides are kept in a humidified chamber. A plastic slide box can be used. Fill the bottom of the box with 1/4 in. of water and place the slides flat, tissue side up, parallel and above the water.

9. Aspirate the proteinase-K solution, using vacuum suction. Wash the tissue at least three times by dropping CMF-DPBS onto each tissue spot with a transfer pipet and aspirating it with vacuum suction. To create a vacuum suction waste container, use a large glass filter flask with vacuum side arms, a vacuum nozzle and two pieces of plastic tubing. Connect the vacuum nozzle to the glass flask with one plastic tube and with the other tube, add a nonfilter pipet tip (200 µl, recommended) to one end and connect the other end to the flask. Use the pipet tip to aspirate reagents from each tissue spot throughout the remaining protocol. When using this method, take care never to touch the tissue.

10. Incubate each spot of tissue with tyramide blocking buffer (according to the manufacturer’s instructions) 60 min at room temperature.

Incubate tissue with primary antibody 11. Make a 1:50 dilution of CD41 primary antibody in tyramide blocking buffer. Remove the tyramide blocking buffer from the tissue using the vacuum suction, add 100 µl diluted CD41 primary antibody to each tissue spot, and incubate 45 to 60 min at room temperature. 12. Remove the primary antibody solution and wash three with 100 µl CMF-DPBS, using vacuum suction to remove the solutions.

Incubate tissue with secondary antibody 13. Make a 1:1000 anti-rat secondary antibody dilution in tyramide blocking buffer, add 100 µl to each tissue spot, and incubate for 30 min at room temperature. 14. Remove the secondary antibody solution and wash three times with 100 µl CMFDPBS, using vacuum suction to remove the solutions.

Amplify signal 15. Add 100 µl of a 1:100 dilution of SA-HRP to each spot and incubate 30 min at room temperature. 16a. If using the ABC-AP Kit in step 20: Add 1 drop of reagent A and 1 drop of reagent B to 5 ml CMF-DPBS/0.1% Tween 20, vortex, and leave on ice at least 30 min before using. The alkaline phosphatase colors Vector Blue and Vector Red work best for CD41 and CD31/Pecam.

16b. If using the ABC Elite Kit in step 20: Add 1 drop of reagent A and 1 drop of reagent B to 2.5 ml CMF-DPBS, vortex, and leave on ice at least 30 min before using. The peroxidase substrate DAB works best for cytokeratin.

17. Remove the SA-HRP solution and wash three times with 100 µl CMF-DPBS, using vacuum suction to remove the solutions. Placental Hematopoietic Stem Cells

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These steps amplify the signal of the primary antibody by detecting horseradish peroxidase (HRP) deposited on the primary/secondary antibody complex. Streptavidin (SA)-HRP will bind to the biotinylated secondary antibodies (which are bound to the primary antibodies).

Amplify signal 18. Add 100 µl tyramide amplification buffer to each spot (according to the manufacturer’s instructions) and incubate 7 to 10 min at room temperature. 19. Remove the tyramide amplification buffer and wash three times with 100 µl CMFDPBS, using vacuum suction to remove the solutions.

Develop substrate 20. Add 100 µl ABC-AP or ABC Elite solution (prepared in step 16) to each tissue spot and incubate 30 min at room temperature. 21. Remove the ABC-AP or ABC Elite solution and wash three times with 100 µl CMF-DPBS, using vacuum suction to remove the solutions. 22. Make the appropriate substrate solution. Add the substrate solution to one slide at a time. Add enough substrate to cover the tissue and watch staining develop under the microscope. Different antibodies and different substrates develop at different speeds. Using a timer to monitor the amount of time it takes one slide to develop can help expedite the procedure for the remaining slides. For example, if CD41 staining requires 5 min. to develop on the first slide, then add the substrate to all remaining slides for only 5 min. and wash. Developing too long can cause increased background or leakiness of the staining. When using alkaline phosphatase substrates add one drop of levamisole for every 5 ml of substrate reagent to block endogenous alkaline phosphatase activity in the tissue. The placenta has high amounts of endogenous alkaline phosphatase.

23. To stop substrate development, aspirate the substrate reagent and wash with 100 µl of the appropriate buffer (according to the substrate manufacturer). Then wash three times with 100 µl CMF-PBS, using vacuum suction to remove the solutions.

Immunostain additional landmark tissue markers 24a. To stain endothelium: Follow steps 11 to 22, but first stain for CD31, using a 1:200 dilution of CD31/Pecam in step 11. Then use a 1:1000 anti-rat secondary antibody dilution in step 13. 24b. To stain trophoblast: Follow steps 11 to 22, but use a 1:1000 dilution of cytokeratin primary antibody in step 11, and omit all tyramide amplification-specific steps (steps 14 and 17). Use normal blocking buffer (5% horse serum/0.05% Tween20/PBS) instead of tyramide blocking buffer in steps 11 to 22. Stain the cytokeratin using a 1:1000 of anti-rabbit secondary antibody dilution in step 13. The second (step 24a) and third (step 24b) stains must include an avidin/biotin blocking kit (used according to the manufacturer’s instructions) because the secondary antibodies are biotinylated.

Mount slides 25. After the last substrate development is complete and the tissues are in PBS, remove all liquid from the top of the slide, using the vacuum suction, one slide at a time. 26. Add at least one drop of mounting medium to each tissue spot, taking care to remove all bubbles. Add a cover slip and let the slide dry for 24 hr. Although in most cases mounting the slides allows for storage for as long as 1 year, it is best to analyze the staining as soon as possible.

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

GO buffer, 10× 3.6 g glucose (Sigma Ultra; 100 mM final) 0.13 g sodium azide (NaN3 , Sigma; 10 mM final) 200 ml water (Cellgro) Store up to 6 months at 4◦ C CAUTION: Sodium azide is toxic! Handle with care.

COMMENTARY Background Information

Placental Hematopoietic Stem Cells

Hematopoietic cells have the ability to selfrenew and generate all cell types of the blood lineages throughout the lifetime of an individual (Weissman, 2000). All HSCs emerge during embryonic development, after which their pool size is maintained by self-renewing cell divisions. As many anatomical sites participate in hematopoiesis during fetal development, resolving the origin of HSCs and the critical developmental events required for their emergence and expansion remains an area of active research. Recently, the mouse placenta was identified as an important fetal hematopoietic organ, responsible for both de novo generation of HSCs and for HSC maintenance and expansion. The placenta becomes a hematopoietic organ shortly after its formation upon fusion of the allantois and chorion (Mikkola et al., 2005). The emergence of HSCs in the placenta starts concomitantly with the aortagonad mesonephrose (AGM) around E9.5 to E10.5, at which time the first hematopoietic progenitors and HSCs, respectively, can be found (Alvarez-Silva et al., 2003; Gekas et al., 2005; Ottersbach and Dzierzak, 2005). At this stage, nascent HSCs/progenitors emerge in the large vessels of the chorionic plate (Rhodes et al., 2008). Subsequently, the number of HSCs in the placenta increases drastically, from ∼2.5, to reaching ultimately ∼50 HSCs at its peak (E12.5-E13.5; Gekas et al., 2005; Ottersbach and Dzierzak, 2005; Rhodes et al., 2008). It is proposed that the placental labyrinth may provide a unique microenvironment that supports the maturation and expansion of HSCs, while protecting them from premature differentiation before the seeding of the fetal liver. Thus, isolation of the placenta enables both the study of a robust population of nascent HSCs and their unique properties, and provides new insights into the specialized

microenvironments in the placenta that support HSCs during embryogenesis. The protocols defined in this unit have been optimized specifically for the mouse placenta and the HSCs it harbors. There are a number of different enzymatic methods used to dissociate organ tissues. We have found that collagenase or collagenase in combination with dispase generates a higher yield of HSCs from the placenta as compared to trypsin (C.G. and H.K.A.M., unpub. observ.; Gekas et al., 2005). Another important factor affecting HSC yield from the placenta is the degree of mechanical dissociation prior to enzymatic treatment. Too little mechanical dissociation prior to enzymatic treatment reduces the accessibility of the tissues for enzymatic activity, whereas forced mechanical dissociation prior to enzymatic dissociation may result in increased cell death and clumping of the cells. The classic phenotypic profile of adult bone marrow HSCs is Lin− Sca-1+ ckit+ CD34− , whereas the phenotype of fetal HSCs changes during their development. The c-kit+ CD34+ profile can be used to enrich for HSCs and multipotential progenitors in all fetal hematopoietic sites, including the placenta (Sanchez et al., 1996; Mikkola et al., 2003; Gekas et al., 2005; Mikkola and Orkin, 2006). However, unlike adult HSCs, nascent fetal HSCs and progenitors co-express CD41, which is downregulated later during maturation of fetal HSCs (Mikkola et al., 2003; Mikkola and Orkin, 2006; Rhodes et al., 2008). In contrast, Sca-1 is not expressed on the surface of nascent HSCs, whereas it becomes upregulated in placental and fetal liver HSCS during their developmental maturation (Mikkola and Orkin, 2006). Of note, Sca-1 is expressed in nonhematopoietic tissues in the placenta (e.g., in placental vasculature and decidua) at a much higher level than in HSCs, and therefore, use of Sca-1 antibody for identification of placental HSCs by flow cytometry

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or immunohistochemistry has to be performed with appropriate costaining and controls. Immunohistochemistry allows identification of the precise cellular niches in which HSCs/progenitors reside. We have observed that immunohistochemistry gives better definition of the placental tissue structure than immunofluorescence, where unstained cells are not visible. However, if colocalization of multiple antigens in the same cell is desired, immunofluorescence with confocal microscopy is recommended. As many of the hematopoietic-specific antigens, e.g., CD41, are not preserved during preparation of paraffin-embedded sections, we routinely use fixed frozen sections for localizing placental

HSCs. Use of tyramide amplification in the staining provides much better signal for many of the hematopoietic antigens.

Critical Parameters and Troubleshooting Dissection It is important to work quickly and to keep embryos and solutions on ice as much as possible in order to maintain cellular viability. A useful indicator is heartbeat, which should be present for at least 30 to 60 min after collection of embryos if they are kept on ice. If placental cells are to be used for culturing or transplantation, it is critical to work under sterile conditions.

Figure 2A.8.5 Localizing hematopoietic stem and progenitor cells in the placental niche. (A) A schematic of the basic structures comprising the placenta. The umbilical cord (the vascular connection between the embryo and placenta) merges with the chorionic plate. The chorionic plate harbors the large vessels of the placenta while the labyrinth contains smaller vessels. The labyrinth is unique in that it contains trophoblast cells, which line the maternal blood spaces and facilitate fetal-maternal exchange. (B) Immunohistochemical stain of a fixed, frozen E10.0 placenta. Nascent HSC/progenitors are marked by CD41 in blue (arrowheads) and are found predominately in the large vessels in the chorionic plate at this time. Endothelial cells are marked by CD31 in red, and trophoblasts are marked by cytokeratin in brown. Notice that at this developmental stage the labyrinth vasculature is just beginning to form and invade within the trophoblast layer (bracket). (C) Immunohistochemical stain of a fixed, frozen E11.5 placenta, where the labyrinth vasculature is more developed. CD41, in blue, marks not only early HSC/progenitors, but also fetal and maternal platelets, which are much smaller in size (∗ ). CD31+ endothelium is red, and cytokeratin+ trophoblasts are brown. Note that most of the CD41+ HSCs/progenitors are in the labyrinth vasculature (arrowheads).

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Single-cell suspension The critical parameters for obtaining highyield single-cell suspensions (also discussed in Background Information) are the degree of mechanical dissociation before the enzymatic action, as well as the incubation time with the enzyme. Too little mechanical dissociation and/or too short an incubation time would result in low cellular yield due to incomplete dissociation; a high level of mechanical dissociation (>20-G needle) and/or very long incubation (>90 min) with collagenase would also result in lower cellular viability and yield. Flow cytometry and FACS sorting It is important to maintain antibody solutions and samples on ice to ensure both preservation of the fluorochromes and high cellular viability, especially for FACS sorting. Immunohistochemistry Overfixation of the tissue can pose a problem for some antigens. Therefore, it is important to fix smaller tissues for less time, ∼2 hr. However, if the tissue is underfixed, the antigen retrieval step can destroy the tissue entirely.

Anticipated Results The protocols above will allow for isolation and visualization of placental tissue and hematopoietic stem and progenitor cells. For a comprehensive summary on expected cellular yield and number of HSCs in placenta and other fetal hematopoietic organs throughout fetal development, see Gekas et al. (2005). For localization of developing HSCs and other hematopoietic cells in the placenta, see Rhodes et al. (2008) and Figures 2A.8.5B and 2A.8.5C.

Time Considerations Timed pregnancies need to be set up 1 to 2 weeks in advance of the experiment, depending on the required embryonic age. Dissection of placentas from one litter (∼10 embryos) should take 1 to 2 hr, depending on dissec-

tion skill and embryonic age. To prepare a single-cell suspension, allow 2 to 2.5 hr. Antibody staining for flow cytometry should take 1 to 2 hr, depending on the number of samples and whether staining with secondary antibodies is required. Preparing placental tissue for immunohistochemistry should take ∼2 days. Likewise, allow 2 days for immunohistochemistry when staining for at least three different markers.

Literature Cited Alvarez-Silva, M., Belo-Diabangouaya, P., Salaun J., and Dieterlen-Lievre, F. 2003. Mouse placenta is a major hematopoietic organ. Development 130:5437-5444. Gekas, C., Dieterlen-Lievre, F., Orkin, S.H., and Mikkola, H.K. 2005. The placenta is a niche for hematopoietic stem cells. Dev. Cell 8:365-375. Mikkola, H.K. and Orkin, S.H. 2006. The journey of developing hematopoietic stem cells. Development 133:3733-3744. Mikkola, H.K., Fujiwara, Y., Schlaeger, T.M., Traver, D., and Orkin, S.H. 2003. Expression of CD41 marks the initiation of definitive hematopoiesis in the mouse embryo. Blood 101:508-516. Mikkola, H.K., Gekas, C., Orkin, S.H., and Dieterlen-Lievre, F. 2005. Placenta as a site for hematopoietic stem cell development. Exp. Hematol. 33:1048-1054. Ottersbach, K. and Dzierzak, E. 2005. The murine placenta contains hematopoietic stem cells within the vascular labyrinth region. Dev. Cell 8:377-387. Rhodes, K.E., Gekas, C., Wang, Y., Lux, C.T., Francis, C.S., Chan, D.N., Conway, S., Orkin, S.H., Yoder, M.C., and Mikkola, H.K.A. 2008. The emergence of hematopoietic stem cells is initiated in the placental vasculature in the absence of circulation. Cell Stem Cell 2:252263. Sanchez, M.J., Holmes, A., Miles, C., and Dzierzak, E. 1996. Characterization of the first definitive hematopoietic stem cells in the AGM and liver of the mouse embryo. Immunity 5:513525. Weissman, I.L. 2000. Stem cells: Units of development, units of regeneration, and units in evolution. Cell 100:157-168.

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Preparation of Hematopoietic Stem and Progenitor Cells from the Human Placenta

UNIT 2A.9

Catherine Robin1 and Elaine Dzierzak1 1

Erasmus MC Stem Cell Institute, Department of Cell Biology, Rotterdam, The Netherlands

ABSTRACT This unit describes a protocol to isolate hematopoietic progenitors and stem cells from human placentae isolated at different time points in development and at the full-term gestational stage. The placenta is extensively washed to eliminate blood contamination on its surface and inside the villi (the vascular compartments of the placenta). The placenta is then mechanically minced into pieces, which are subsequently digested with an enzyme cocktail. After dissociation and Þltration, placental cells are available for further phenotypic and functional analyses. Curr. Protoc. Stem Cell Biol. 14:2A.9.1C 2010 by John Wiley & Sons, Inc. 2A.9.8.  Keywords: human placenta r enzymatic treatment r hematopoietic stem and progenitor cell isolation

INTRODUCTION This unit describes a protocol to mechanically dissociate human placentae collected at different time points during development (between 3 and 19 weeks), including full term (Basic Protocol 1). The placenta tissue is then further dissociated by enzymatic treatment to obtain a single-cell suspension (Basic Protocol 2). Both procedures are designed to obtain the most efÞcient hematopoietic cell recovery in terms of number and viability. Placenta cells can be frozen and stored, or used immediately after isolation. The in vivo and in vitro hematopoietic potential of placenta cells can be subsequently studied for the presence of hematopoietic stem cells and progenitors, respectively (Robin et al., 2009). NOTE: The entire procedure is performed in a laminar-ßow hood with sterile medium and materials. All materials coming into contact with live placental cells must be sterilized. NOTE: All incubations are performed in a humidiÞed 37◦ C, 5% CO2 incubator. NOTE: Media and solutions used to wash the placenta and collect the cells are kept cold. Medium for the enzymatic steps is prewarmed at 37◦ C before use.

MECHANICAL DISSOCIATION OF HUMAN PLACENTA The outside of the placenta (Fig. 2A.9.1A, maternal side; Fig. 2A.9.1B, embryonic side) is extensively washed to eliminate all blood clumps attached to it. The blood in the villi is ßushed away by extensive and repeated injection of medium into the vein and arteries of the placental cord. The procedure is not fully applicable to early developmental stage placentae, since the integrity of the tissue is usually compromised. The washed placenta is subsequently cut into small pieces and prepared for the enzymatic digestion.

BASIC PROTOCOL 1

Materials Human placenta PBS supplemented with EDTA (PBS/EDTA; see recipe) PBS supplemented with 10% (v/v) fetal bovine serum (PBS/FBS; see recipe) Current Protocols in Stem Cell Biology 2A.9.1-2A.9.8 Published online August 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02a09s14 C 2010 John Wiley & Sons, Inc. Copyright 

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A

cotyledons

connective tissue septa

B umbilical cord

placenta vasculature

clamp

Figure 2A.9.1 Full-term human placenta obtained after birth. (A) Maternal aspect. The side shown faces the uterine wall. (B) Fetal aspect. The side shown faces the baby with the umbilical cord on the top. The white fringes surrounding the placenta are the remnants of the amniotic sac.

Ficoll Collagenase (use at 0.125% (w/v) after dilution 1/20 in PBS/FBS; see recipe)

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50-ml collection tubes 10-ml plastic pipet Absorbent paper Large stainless steel trays (to hold the placenta and ßuids during wash procedure) 50-ml syringe 18-G needles Clamp Large glass or plastic petri dishes (20-cm diameter) Cutting board Carving knives Forceps Current Protocols in Stem Cell Biology

Collect and prepare cord blood The cord blood is a well known source of hematopoietic stem/progenitor cells. It is used as a quantitative and qualitative control for comparison of hematopoietic stem/progenitor cells obtained from other tissues, such as the placenta. 1. Collect cord blood in one or more 50-ml collection tubes, each containing 10 ml of PBS/EDTA 2. Dilute the cord blood 1:2 (v/v) into PBS/FBS. 3. Place 20 ml diluted cord blood cells on the top of 20 ml Ficoll in a 50-ml centrifuge tube for density gradient fractionation. 4. Centrifuge 20 min at 670 × g, room temperature, with low deceleration so as not to perturb the mononuclear cell ring. 5. Aspirate the mononuclear cell ring with a 10-ml plastic pipet. 6. Wash cells twice in 50 ml cold PBS/FBS, centrifuging 10 min at 170 × g, 4◦ C, each time. Resuspend the cells in 1 ml of PBS/FBS. 7. Store cells for several hours at 4◦ C until further use.

Collect placenta 8. Prepare the surface of a laminar-ßow hood by covering it with absorbent paper in case of blood spillage. 9. Collect placentae into PBS/EDTA. Early stage human placentae are obtained from elective abortions. Gestational age is determined by ultrasound fetal measurements. Term placentae are obtained either by cesarean section or from vaginal deliveries. All placentae are obtained with informed consent. The placenta is collected in a small plastic bucket and PBS/EDTA is added to cover it completely. Placentae can be used directly or after overnight storage at 4◦ C.

10. In a stainless steel tray, wash the outside of the placenta extensively with cold PBS/EDTA to eliminate all dead tissues and blood clumps. 11. Remove the amniotic and deciduas membranes, and cut all but the proximal 10 cm of the umbilical cord. 10 cm of cord are kept attached to the placenta to allow the placement of a clamp.

Collect placental blood 12. Unclamp the umbilical cord, hold the fetal side of the placenta downward, and place the umbilical cord over a 50-ml tube. Collect the blood from inside the placenta vasculature by manually squeezing the placenta to allow evacuation of the blood through the umbilical cord. Aspirate the remaining blood through an 18-G needle attached to a 50-ml syringe. The collected placental blood is similar to umbilical cord blood.

13. Extensively wash the villi by repeated injections of 50 ml cold PBS/EDTA through an 18-G needle attached to a 50-ml syringe. Flush the PBS away by manually squeezing the placenta (fetal side down), with the umbilical cord section placed in a 50-ml tube. The injection of PBS is done via the vein and arteries of the cord. If there are blood clumps in the cord, the injection can be done directly into the large vessels of the vascular labyrinth (this procedure is incompatible with step 6). The procedure must be repeated at least 10 times, until the placental vasculature appears white.

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14. To collect the cells attached inside the placenta vasculature, inject via the cord vessels 20 to 50 ml prewarmed PBS/FBS containing collagenase (0.125% w/v) through an 18-G needle attached to a 50-ml syringe. Inject the volume of medium needed to Þll most of the vessels.

15. Clamp the cord and place the placenta in a large petri dish. Incubate the placenta for 1 hr in an incubator at 37◦ C. 16. Collect intravascular cells detached by the collagenase treatment by aspiration via the cord vessels in a 50-ml tube. 17. Wash the cells collected after collagenase treatment twice, each time in 50 ml of cold PBS/FBS, centrifuging 10 min at 170 × g, 4◦ C, each time.

Mince the tissue 18. Place placenta tissue on a cutting board and mince into small pieces with a carving knife and forceps. Do not allow the tissue and pieces to dry; if necessary, add PBS/FBS. Placenta tissues are very difÞcult to cut with scissors—the authors suggest using highly sharpened carving knifes.

19. In preparation for the enzymatic digestion, place the placenta pieces into a 50-ml tube. BASIC PROTOCOL 2

ENZYMATIC DISSOCIATION OF HUMAN PLACENTA The placenta is a large, dense, highly vascular tissue that is difÞcult to dissociate. Several protocols for enzymatic dissociation of the placenta have been tested, and the authors present here a protocol that is, to date, the most efÞcient processing procedure leading to the recovery of the highest number of viable hematopoietic cells.

Materials Placenta pieces (Basic Protocol 1) Enzyme cocktail (see Table 2A.9.1 PBS/FBS (see recipe) Ficoll 50-ml tubes ParaÞlm 37◦ C water bath with shaking 10-ml plastic pipet Sterile cotton gauze placed in a stainless steel soup strainer on top of a sterile glass beaker (500 ml) Cell strainer (40-μm Nylon) NOTE: Keep all cells at 4◦ C and carry out all procedures except Ficoll separation at 4◦ C. Table 2A.9.1 Preparation of the Enzymatic Cocktail for the Digestion of Placenta Pieces

Volume of each reagent (ml)a Preparation of Hematopoietic Stem and Progenitor Cells from the Human Placenta

Tissue (g) Collagenase 50

10

Pancreatin

Dispase

DNase I

24

13.3

2

PBS/FBS Final volume 150.7

200

a See the Reagents and Solutions section for the solution recipes.

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Treat placenta pieces with enzymes 1. Place 10 to 15 g of placenta pieces into a 50-ml tube. To dissolve a larger placenta portion, place the placenta pieces into a glass bottle (500 ml) instead of the 50-ml tube. Up to 15 g of placenta pieces can be digested in a Þnal volume of 50 ml. However, the authors have established that 5 g of placenta tissue in 200 ml Þnal volume gives optimal enzymatic digestion.

2. Fill the tube with the enzyme cocktail (Table 2A.9.1) The enzymatic treatment is performed in the presence of 0.001 mg/ml DNase. Cell death occurs during the procedure, and DNA from these cells must be digested, or the cell pellet will be inseparable from the viscous solution containing high-molecular-weight DNA strands.

3. Gently mix the tubes and seal with ParaÞlm. 4. Incubate the tubes in a water bath at 37◦ C for 1 to 1.5 hr under agitation. The authors found that incubation longer than 1.5 hr does not improve the dissociation much but it does increase cell death.

5. Dissociate tissues further by repeated pipetting with a 10-ml plastic pipet. This procedure can be difÞcult in the presence of large placenta pieces that did not dissociate.

6. Pass the cell suspension through sterile cotton gauze placed in a stainless steel soup strainer on top of a sterile glass beaker (500 ml). Passing the cell suspension through the gauze helps to eliminate all nondigested tissue clumps and debris.

Wash the Þltrate 7. Wash the Þltrate with 25 ml PBS/FBS and remove the gauze without squeezing. Replace the gauze after Þltration of every ∼50 ml of suspension.

8. Dilute the placenta suspension 1:2 with cold PBS/FBS. 9. Centrifuge 10 min at 170 × g, 4◦ C. 10. Remove the supernatant and wash the cell pellet twice with 50 ml of PBS/FBS. 11. Resuspend the cells in 20 ml of PBS/FBS and place the cell suspension on top of 20 ml Ficoll in a 50-ml tube for density gradient fractionation. 12. Centrifuge 20 min at 670 × g, room temperature, with low deceleration to keep the mononuclear cell ring unperturbed.

Collect the cells 13. Collect the mononuclear cell ring with a 10-ml plastic pipet and place into a 50-ml tube. 14. Wash the cells collected after Ficoll treatment twice, each time in 50 ml of cold PBS/FBS, centrifuging 10 min at 170 × g, 4◦ C, each time. 15. Filter cells through a 40-μm nylon cell strainer. Cells can be kept at 4◦ C for several hours, until use in hematopoietic assays or in preparation for frozen storage at –80◦ C. After Ficoll gradient enrichment, the placenta cell suspension contains mononuclear cells. The cell suspension contains hematopoietic progenitors (as tested by in vitro clonogenic assay and ßow cytometry analyses) and hematopoietic stem cells (as tested by in vivo transplantation into immunodeÞcient mice; Robin et al., 2009). However, cell clump formation often occurs. The authors recommend reÞltering the cell suspension before any use (e.g., freezing, ßow cytometry analysis/sort). Current Protocols in Stem Cell Biology

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Dispase (neutral protease grade I) stock solution (5 mg/ml) Dissolve the dispase in sterile Milli-Q-puriÞed water (5 mg into 1 ml). The lyophilized enzyme is stable at 2◦ to 8◦ C until the expiration date printed on the label. The solution is stable at −15◦ to −25◦ C until the expiration date printed on the label. Be careful when opening the vial to avoid the powder bursting out (if it is kept under vacuum). Strive to dissolve all powder (including what is attached to the cap) to get the correct concentration.

DNase stock solution (1:100) Dissolve DNase I in 1 ml of sterile Milli-Q-puriÞed water and transfer to a 50-ml tube Add 20 ml of sterile Milli-Q-puriÞed water into the tube and mix well When stored at −20◦ C, the enzyme is stable through the expiration date printed on the label.

Pancreatin stock solution (2.5% w/v) Prepare a 0.5% (w/v) PVP solution (polyvinylpyrolidone K30; Fluka) by adding 2.5 g of PVP powder in 500 ml of sterile phosphate-buffered saline. Shake the tube vigorously to completely dissolve the powder and obtain a clear solution. Dissolve 5 g of pancreatin powder (pancreatin from porcine pancreas) in 200 ml of 0.5% PVP solution, add a sterile magnetic stirrer, and agitate 30 min at 4◦ C. After 30 min, the solution remains cloudy.

Aliquot 1.5 ml of solution per 1.5-ml microcentrifuge tube and centrifuge 30 min at 15,000 × g, 4◦ C. Pool all supernatant in 50-ml tubes and discard the pellets. The solution is stable at −20◦ C until the expiration date printed on the label.

PBS/EDTA Phosphate-buffered saline (PBS) supplemented with EDTA, penicillin (100 U/ml), and streptomycin (100 mg/ml). Store up to 1 month at 4◦ C. Add 1.5 mg of EDTA per milliliter of cord blood or placenta cell suspension. This solution is used for placenta and blood collection and wash medium.

PBS/FBS Phosphate-buffered saline (PBS) supplemented with 10% (v/v) fetal bovine serum (FBS), penicillin (100 U/ml) and streptomycin (100 mg/ml). Store up to 1 month at 4◦ C. This solution is used as placenta and blood cell resuspension medium. Preparation of Hematopoietic Stem and Progenitor Cells from the Human Placenta

Type I collagenase stock solution (2.5% w/v) Dissolve 2.5 g of collagenase powder in 100 ml of sterile PBS. Filter the solution using a 0.2-μm Þlter and divide into 50-ml aliquots. Store up to several months at −20◦ C without repeated freeze-thaws.

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COMMENTARY Background Information The placenta plays a crucial role during embryonic and fetal development. It connects the developing fetus to the uterine wall and allows nutrient uptake, waste elimination, and gas exchange between the mother and the developing fetus (Gude et al., 2004). During human embryonic development, hematopoietic stem cells (HSCs) and progenitors are found in different anatomical sites that share the particular feature of being highly vascularized (Tavian et al., 2001). The Þrst hematopoietic activity is detected in the yolk sac, starting at day 16 of development, with the production of differentiated erythrocytes. Later on, hematopoiesis takes place also in the embryo: in the aortagonad-mesonephros (AGM) region, umbilical and vitelline arteries, and fetal liver. The spatio-temporal hematopoietic events observed in humans follow what has been previously found in the mouse embryo model (Tavian and Peault, 2005). Another highly vascularized tissue, the placenta, was recently reported as a potent hematopoietic site during both mouse (Alvarez-Silva et al., 2003; Gekas et al., 2005; Ottersbach and Dzierzak, 2005; Ziegler et al., 2006; Corbel et al., 2007) and human (Barcena et al., 2009a,b; Robin et al., 2009) embryonic development. Around day 24 of human development, primitive erythroblasts Þll the placental vasculature (Challier et al., 2005). Multipotent progenitors and HSCs (deÞned by their ability to multilineage repopulate irradiated NOD-SCID recipients) start to be detected as early as week 6 in gestation and are present through to term (Robin et al., 2009). In the mouse embryo, the chorion and allantois (the early embryonic tissues that fuse to form the placenta) can both generate and support hematopoietic progenitor cells before the circulation is established between embryo and placenta, as shown by in vitro clonogenic assay (Ziegler et al., 2006; Corbel et al., 2007). At mid-gestation, the placenta contains more HSCs and progenitors than the AGM and yolk sac (Gekas et al., 2005). However, it is as yet uncertain if the placenta can generate de novo HSCs (Rhodes et al., 2008). Thus, the placenta provides a suitable microenvironment or niche throughout development and until term for HSC maintenance and ampliÞcation, similar to the fetal liver. Based on this observation, many cell lines have been isolated from human placentae at a wide range of developmental stages to test if they constitute a suitable feeder to maintain/expand hematopoietic cell populations (Miyamoto et al., 2004; Current Protocols in Stem Cell Biology

Zhang et al., 2004; Kim et al., 2007; Robin et al., 2009). The protocol described in this review was speciÞcally developed to isolate hematopoietic stem and progenitor cells with an optimal viability and yield.

Critical Parameters and Troubleshooting The procedure of placenta dissociation generates a high degree of cell mortality. To improve the procedure to harvest viable cells, DNase I must be added during the enzymatic digestion. Cells must be kept at 4◦ C after isolation and be processed as soon as possible for storage or functional testing. A complete digestion of the placenta (particularly at later gestational stages or term) is very difÞcult due to the large vessels and the tight adhesive cells that form the villi. The authors tested several enzymatic procedures, with different enzymes at different concentrations, and found that the combination pancreatin/dispase/collagenase was the most efÞcient to isolate hematopoietic cells with the best viability and yield. The washing and Þltering steps are crucial and must be performed rigorously.

Anticipated Results This protocol generates large numbers of mononucleated hematopoietic cells after enzymatic treatment and Ficoll separation. For fullterm placenta (average = 460 g), the number of cells averages at 240 × 106 mononucleated cells/placenta.

Time Considerations The dissociation of a complete full-term placenta takes a complete day and at least two research personnel. This has to be taken into consideration for further analysis of the cells. The authors recommend freezing and storing the cells just after isolation and performing further analyses on another day.

Acknowledgements We thank all former and current laboratory members who contributed to the elaboration of the placenta dissociation protocol. We are also grateful to the tissue donors for their pivotal contributions to the study of the human placenta.

Literature Cited Alvarez-Silva, M., Belo-Diabangouaya, P., Salaun, J., and Dieterlen-Lievre, F. 2003. Mouse placenta is a major hematopoietic organ. Development 130:5437-5444.

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Barcena, A., Kapidzic, M., Muench, M.O., Gormley, M., Scott, M.A., Weier, J.F., Ferlatte, C., and Fisher, S.J. 2009a. The human placenta is a hematopoietic organ during the embryonic and fetal periods of development. Dev. Biol. 327:2433. Barcena, A., Muench, M.O., Kapidzic, M., and Fisher, S.J. 2009b. A new role for the human placenta as a hematopoietic site throughout gestation. Reprod. Sci. 16:178-187. Challier, J.C., Dubernard., G., Galtier, M., Bintein, T., Vervelle, C., Raison, D., Espi´e, M.J., and Uzan, S. 2005. Immunocytological evidence for hematopoiesis in the early human placenta. Placenta 26:282-288. Corbel, C., Salaun, J., Belo-Diabangouaya, P., and Dieterlen-Lievre, F. 2007. Hematopoietic potential of the pre-fusion allantois. Devel. Biol. 301:478-488. Gekas, C., Dieterlen-Lievre, F., Orkin, S.H., and Mikkola, H.K. 2005. The placenta is a niche for hematopoietic stem cells. Dev. Cell 8:365375. Gude, N.M., Roberts, C.T., Kalionis, B., and King, R.G. 2004. Growth and function of the normal human placenta. Thromb. Res. 114:397407. Kim, S.J., Song, J.H., Sung, H.J., Yoo, Y.D., Geum, D.H., Park, S.H., Yoo, J.H., Oh, J.H., Shin, H.J., Kim, S.H., Kim, J.S., and Kim, B.S. 2007. Human placenta-derived feeders support prolonged undifferentiated propagation of a human embryonic stem cell line, SNUhES3: Comparison with human bone marrow-derived feeders. Stem Cells Dev. 16:421-428. Miyamoto, K., Hayashi, K., Suzuki, T., Ichihara, S., Yamada, T., Kao, Y., Yamabe, T., and Ito, Y. 2004. Human placenta feeder layers support undifferentiated growth of primate embryonic stem cells. Stem Cells 22:433-440.

Ottersbach, K. and Dzierzak, E. 2005. The murine placenta contains hematopoietic stem cells within the vascular labyrinth region. Dev. Cell 8:377-387. Rhodes, K.E., Gekas, C., Wang, Y., Lux, C.T., Francis, C.S., Chan, D.N., Conway, S., Orkin, S.H., Yoder, M.C., and Mikkola, H.K. 2008. The emergence of hematopoietic stem cells is initiated in the placental vasculature in the absence of circulation. Cell Stem Cell 2:252-263. Robin, C., Bollerot, K., Mendes, S., Haak, E., Crisan, M., Cerisoli, F., Lauw, I., Kaimakis, P., Jorna, R., Vermeulin, M., Kayser, M., van der Linden, R., Imanirad, V., Verstegen, M., Nawaz-Jousef, H., Papazian, N., Steegers, E., Cupedo, T., and Dzierzak, E. 2009. Human placenta is a potent hematopoietic niche containing hematopoietic stem and progenitor cells throughout development. Cell Stem Cell 5:385395. Tavian, M. and Peault, B. 2005. Embryonic development of the human hematopoietic system. Int. J. Devel. Biol. 49:243-250. Tavian, M., Robin, C., Coulombel, L., and Peault, B. 2001. The human embryo, but not its yolk sac, generates lympho-myeloid stem cells: mapping multipotent hematopoietic cell fate in intraembryonic mesoderm. Immunity 15:487-495. Zeigler, B.M., Sugiyama, D., Chen, M., Guo, Y., Downs, K.M., and Speck, N.A. 2006. The allantois and chorion, when isolated before circulation or chorio-allantoic fusion, have hematopoietic potential. Development 133:41834192. Zhang, Y., Li, C., Jiang, X., Zhang, S., Wu, Y., Liu, B., Tang, P., and Mao, N. 2004. Human placenta-derived mesenchymal progenitor cells support culture expansion of long-term cultureinitiating cells from cord blood CD34+ cells. Exp. Hematol. 32:657-664.

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Isolation and Characterization of Mesoangioblasts from Mouse, Dog, and Human Tissues

UNIT 2B.1

Rossana Tonlorenzi,1 Arianna Dellavalle,1 Esther Schnapp,1 Giulio Cossu,1 and Maurilio Sampaolesi1 1

Stem Cell Research Institute, San Raffaele Scientific Institute, Milan, Italy

ABSTRACT Mesoangioblasts are recently identified stem/progenitor cells, associated with small vessels of the mesoderm in mammals. Originally described in the mouse embryonic dorsal aorta, similar though not identical cells have been later identified and characterized from postnatal small vessels of skeletal muscle and heart (not described in this unit). They have in common the anatomical location, the expression of endothelial and/or pericyte markers, the ability to proliferate in culture, and the ability to undergo differentiation into various types of mesoderm cells upon proper culture conditions. Currently, the developmental origin of mesoangioblasts, their phenotypic heterogeneity, and the relationship with other mesoderm stem cells are not understood in detail and are the subject of active research. However, from a practical point of view, these cells have been successfully used in cell transplantation protocols that have yielded a significant rescue of structure and function in skeletal muscle of dystrophic mice and dogs. Since the corresponding human cells have been recently isolated and characterized, a clinical trial with these cells is planned in the near future. This unit provides detailed methods for isolation, culture, and characterization of mesoangioblasts. Curr. Protoc. Stem Cell Biol. 3:2B.1.1-2B.1.29. C 2007 by John Wiley & Sons, Inc.
Keywords: mesoangioblasts r pericytes r mesoderm progenitor cells r cell culture

INTRODUCTION The protocols in this unit are designed to provide a basis for the isolation, cloning, and propagation of mesoangioblasts derived from mouse embryo aorta (see Basic Protocol 1), adult mouse skeletal muscle (see Basic Protocol 2), human adult skeletal muscle (see Alternate Protocol 1), and dog adult skeletal muscle (see Alternate Protocol 2). Various differentiation methods are also described: co-culture with C2C12 myoblasts (see Basic Protocol 3), co-culture with rat L6 myoblasts (see Alternate Protocol 3), spontaneous differentiation (see Alternate Protocol 4), induction of smooth muscle with TGF (see Alternate Protocol 5), induction of osteoblasts with BMP2 (see Alternate Protocol 6), and induction of adipocytes (see Alternate Protocol 7). In addition, this unit describes inactivation of STO cells or mouse embryo fibroblasts (MEF) by mitomycin C for use as a feeder layer (see Support Protocol 1), collagen (see Support Protocol 2) and matrigel (see Support Protocol 3) coating of tissue culture surfaces, and freezing procedures for mesoangioblasts and pericyte-dervied cells (see Support Protocol 4). Successful derivation and propagation of mesoangioblasts require basic animal handling, dissection, and tissue culture skills. Characterization requires basic histochemistry, biochemistry, and molecular biology skills. NOTE: Mesoangioblasts and pericyte-derived cells must be cultured under physiological O2 conditions (5% O2 , 5% CO2 , 90% N2 ; see Critical Parameters). Somatic Stem Cells Current Protocols in Stem Cell Biology 2B.1.1-2B.1.29 Published online December 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02b01s3 C 2007 John Wiley & Sons, Inc. Copyright


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NOTE: All procedures described in this unit should be performed under sterile conditions in either Class II biohazard flow hoods or laminar flow horizontal draft hoods. When working with human material, Class II biohazard flow hoods are recommended. NOTE: The protocol should be approved by the Institutional Animal Care and Use Committee (IACUC), even though these procedures do not cause any suffering to the animals employed—in the case of dogs, a muscle biopsy is performed under local anesthesia; in the case of human material, approval of the Institutional Ethics Committee and informed consent from the patients are required. BASIC PROTOCOL 1

ISOLATION, CLONING, AND PROPAGATION OF MESOANGIOBLASTS FROM MOUSE EMBRYONIC AORTA Primary culture of tissue fragments from mouse embryonic aorta results in the outgrowth from the explant of a mixed population of cells that includes mesoangioblasts. The progressive increase in the proportion of mesoangioblasts through this outgrowth phase (due to the inability of many other cell types to proliferate in vitro under these conditions) allows for their isolation and efficient cloning.

Materials Dissected aorta (three mouse embryos at embryonic day 10.5) D20 medium (see recipe), sterile 3.5-cm collagen-coated petri dishes (see Support Protocol 2) Phosphate-buffered saline without Ca2+ /Mg2+ (CMF-PBS; Sigma cat. no. D8537), sterile Collagenase/dispase solution (see recipe, Sigma), sterile Fetal bovine serum (heat-inactivated FBS; Cambrex), sterile Trypan blue (Sigma cat. no. T8154) 48-well plates (Nunc) coated with mitotically inactivated STO cells (see Support Protocol 1) 0.05%/0.02% (w/v) trypsin/EDTA (Sigma cat. no. T3924), sterile Curved and straight forceps, sterile Rounded-edge disposable scalpels, sterile 3.5-, 6-, and 15-cm petri dishes 1-ml sterile syringes and insulin needles 37◦ C, 5% CO2 /5% O2 /90% N2 humidified (water-saturated) incubator 15-ml centrifuge tubes 37◦ C water bath Hemacytometer Dissection microscope 48-well plates 25- and 75-cm2 vented tissue culture flasks (Nunc) Additional reagents and equipment for trypan blue exclusion (UNIT 1C.3) Dissect aorta-gonads-mesonephrons 1. Carefully collect the embryos at embryonic day 10.5 (E 10.5) in D20 medium.

Isolation and Characterization of Mesangioblasts

E 11.5 is equally fine, but at later stages, the anatomy becomes more complex and dissection becomes more difficult. At earlier stages, dissection is also more difficult as the two aortas are still separated and closely adherent to the lateral side of the paraxial mesoderm, making contamination from somitic cells more likely. Earlier stages should be avoided unless necessary (e.g., when all or most mutant embryos die or are already severely abnormal at E 10).

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2. Holding the embryo flat on its back with straight forceps at the thoracic level, eviscerate it with curved forceps. Once the intestine is removed the aorta-gonadsmesonephrons (AGM) becomes visible. The AGM can be easily distinguished because the vessels contain blood and are surrounded on both sides by the segmented mesonephrons.

3. Make a transverse cut above and below the AGM with a rounded-edge scalpel and gently remove the AGM with curved forceps. For further details, consult Hogan et al. (1994).

4. Transfer dissected AGM into a new 6-cm petri dish containing 5-ml D20 medium. Make all transfers of tissue in the liquid drop that forms between the adjacent but not tightly closed edges of the curved forceps used for dissection (see also step 8). Do not use glass or plastic pipets because tissues tend to attach to glass and plastic.

5. Holding the aorta with a 1-ml sterile syringe and insulin needle, dissect away the mesonephrons with two cuts parallel to the longitudinal axis of the aorta. 6. Sharply cut the isolated vessels into 1- to 2-mm size fragments using sterile insulin needles. Proceed immediately to establish cultures (aortic fragments cannot be stored). Start each culture with no less than five or six fragments per 3.5-cm petri dish, since a certain density is necessary for the initial outgrowth of mesoangioblasts.

Initiate primary cultures 7. Pre-treat the appropriate number of 3.5-cm collagen-coated petri dishes by pipeting 1.5 ml D20 medium into each dish, making sure that the surface is completely covered. Gently aspirate the medium, but not completely so that the dish surface remains thoroughly wet. 8. Carefully transfer the aortic fragments (up to ten) into each 3.5-cm pre-treated collagen-coated dish. Avoid aspirating the fragments with any plastic tip or glass Pasteur pipet because the tissue is very sticky and may adhere to their internal surface.

9. After the fragments have been transferred into dishes, add 700 µl D20 medium by slowly pipetting it along the edge of the dish to prevent detachment and floating of fragments. 10. Create a humidified chamber for the cultures by placing up to six 3.5-cm dishes containing aortic fragments into a 15-cm dish that also contains a 3.5-cm dish without cover and filled with sterile distilled water. Because an increase in salt concentration due to medium evaporation in the incubator, may be lethal for the cells, it is necessary to culture the fragments in a humidified chamber.

11. Place cultures overnight in a 37◦ C, 5%CO2 /5% O2 /90% N2 incubator. 12. Approximately 24 hr after assembly of cultures, carefully add 1 ml D20 medium to each dish. Return cultures to the incubator. At this time, if the aorta fragments have been manipulated properly, initial outgrowth of adherent cells (mainly large, flat fibroblasts) should be apparent under microscope inspection within 24 hr. After 3 to 7 days, mesoangioblasts should start to be distinguishable as small, round, very refractile cells, weakly adhering to the underlying flat cells (Fig. 2B.1.1A). Somatic Stem Cells

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Figure 2B.1.1 Morphology and immunocytochemistry of mesoangioblasts. (A) Typical outgrowth of small, refractile, poorly adhering mesoangioblasts from an explant of mouse E 10.5 dorsal aorta (phase contrast). (B) An emerging clone of human pericyte-derived cell: absence of a feeder layer makes it easier to appreciate the morphology of the colony (phase contrast). (C) Proliferating mouse embryonic mesoangioblasts (phase contrast). (D) Proliferating mouse adult mesoangioblasts from skeletal muscle (phase contrast). (E) Proliferating dog adult mesoangioblasts from skeletal muscle (phase contrast). (F) Proliferating human adult pericyte-derived cells from skeletal muscle (phase contrast). (G) Senescent human adult pericyte-derived cells from skeletal muscle at 25 PD (population doublings). Note flat, large adhering cells that rarely divide (phase contrast). (H) Mutlinucleated myotubes developed in vitro from human pericyte-derived cells from skeletal muscle (phase contrast). (J) Immunofluorescence of mouse embryonic mesoangioblasts treated with TGF beta and then stained with an antibody recognizing smooth alpha actin (red). Nuclei are stained with DAPI (blue). (K,L) Alkaline phosphatase staining of mouse embryonic (K) and mouse adult (L) mesoangioblasts. Alkaline phosphatase staining of the same cells after exposure to BMP2 as shown in O and P, respectively. (N) Oil-Red O staining of lipid droplets of mouse embryonic mesoangiobalsts induced to adipose differentiation. (M,Q) Myogenic differentiation of human pericyte-derived cells in co-culture with rat L6 myoblasts: myotubes are stained red by anti-sarcomeric myosin antibody; human nuclei are stained green by the anti-lamin A/C antibody (M) while all nuclei are stained blue by DAPI (Q). Magnifications: 200×: A, C-E, H, K, L, O, P; 400×: B, F, G, J, M, N, Q.

Dissociate primary mouse embryo aorta culture 13. Remove D20 culture medium from the dish and rinse two times with 1 ml CMFPBS at room temperature, each time. After the second rinse, remove CMF-PBS completely. Isolation and Characterization of Mesangioblasts

Carefully aspirate and pipet liquids on one side, tipping the dish and avoiding touching either the aorta fragments or surrounding cells.

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14. Add 2 ml collagenase/dispase solution to each dish, and remove cells first and then the aortic fragments by gentle pipetting and mild scraping using a pipettor. Slight adjustments in enzyme concentration and/or digestion time (usually 5 to 15 min) may be necessary depending upon the batch of enzymes used.

15. Transfer the cell and tissue suspension into a 15-ml centrifuge tube. 16. Repeat steps 14 and 15 three additional times. 17. Add an additional 2 ml of collagenase/dispase solution directly to the centrifuge tube. The digestion of tissue fragments and cells is performed in a final volume of 10 ml for each dish.

18. Incubate 15 min in a 37◦ C water bath. Flick and invert the tube three times during incubation, monitoring the dissociation of tissue.

19. Stop the reaction by adding 3 ml FBS to the tube. Centrifuge 15 min at 232 × g, room temperature. 20. Discard supernatant and resuspend the pellet in 200 µl of D20 medium. Pipet up and down several times using a pipet with a filtered tip to disaggregate any residual tissue clumps. A correct digestion should result in an almost homogeneous suspension of cells. Allow sedimentation of small clumps and undigested fragments. Collect supernatant.

21. Count viable cells by trypan blue exclusion (UNIT 1C.3) using a hemacytometer. Proceed immediately to cloning. In parallel to cloning, a small aliquot (30 µl) of total cell suspension should be plated in a single well of a 48-well collagen-coated plate to check cell survival rate.

Clone mouse embryo aorta mesoangioblasts 22. Dilute cells in D20 medium to obtain 150 ml of each of the following concentrations: 1 cell/ml 10 cells/ml 20 cells/ml 30 cells/ml. A total volume of ∼150 ml of cell suspension is needed for three 48-well plates.

23. Aspirate the medium from the 48-well plates coated with mitotically inactivated STO. To avoid the risk of drying the feeder layer, aspirate the medium from no more than three plates at a time.

24. For each concentration, plate 1 ml/well in three 48-well plates. As a control, keep two 48-well plates of inactivated feeder layer STO in 1 ml D20 medium/well without the addition of any cell suspension for at least 1 month. The eventual proliferation of MMC-resistant (or incompletely inactivated) cells should be periodically monitored under the microscope.

25. Prepare a humidified chamber by placing the 48-well plates into a clean plastic box, along with two open 6-cm petri dishes filled with sterile distilled water. Cover the box with aluminum foil.

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26. Place cultures in a 37◦ C, 5% CO2 /5% O2 /90% N2 humidified incubator for at least 1 week. A dedicated incubator should be used, or at least a rarely opened incubator.

27. After 1 week, carefully inspect the cultures with a microscope to distinguish the first clones (see Fig. 2B.1.1B). If clones appear in dishes plated with 1 cell/well, discard dishes plated at higher density. 28. Add 200 µl D20 medium to each well. 29. Passage the clones when the cells have covered ≥50% of the well surface. If cells are healthy and growing properly, a clear acidification of the medium (color turning to orange) should be evident.

Sub-culture mouse embryo aorta mesoangioblasts 30. At the time of first passage, carefully aspirate the medium and rinse each well with 1 ml of CMF-PBS at room temperature. 31. Add 200 µl of 0.025% trypsin/EDTA to each well. Incubate 5 to 10 min at 37◦ C, monitoring under microscope for complete detachment of cells. 32. Inactivate trypsin by adding 800 µl D20 medium down the growing surface of each well. Carefully collect all cells. 33. Transfer cells and medium to a 15-ml centrifuge tube and centrifuge 5 min at 232 × g, room temperature. 34. Discard supernatant, suspend the pellet in 1 ml fresh D20 medium and plate in new, uncoated well of a 48-well plate without the feeder layer. After the first passage to uncoated plastics, mesoangioblasts will loose their round, refractile appearance and will acquire a new morphology of small, triangular, adherent cells (see Fig. 2B.1.1C), which they will maintain until senescence (characterized by a large, flat morphology as shown in Fig. 2B.1.1G for similarly appearing human senescent cells). From this step on, no more feeder layer will be necessary, but particular attention will have to be paid to the density of cells. Until the third/fourth passage, cells must be grown at high density and must be split when fully confluent into progressively larger wells (from 48- to 24- to 12- to 6-well plates and later to 25- and 75-cm2 tissue culture flasks). This phase is the most critical for mesoangioblast derivation; in fact, many clones may differentiate or go to senescence and/or stasis; if culture conditions are inadequate, all clones may be lost at this stage. The successful, continuously proliferating clones usually represent a small percentage of all subcultured clones (∼5% to 10%). A clone can be considered “established” if cells proliferate at a regular rate (∼12 hr doubling time), maintain a typical morphology (see Fig. 2B.1.1C). Once established, all clones (or at least a significant number ≥10) need to be propagated and characterized (see Basic Protocol 3 and Alternate Protocols 3 to 7).

Propagate and freeze established mouse embryo aorta mesoangioblast clones Once established and expanded, mesoangioblast clones can be maintained in the absence of feeder layer and grown in 25- and 75-cm2 vented tissue culture flasks. Split the cells when 70% to 80% confluent, at split ratios up to 1:4. Change the medium every 3 days. 35. Aspirate and discard the medium, and rinse with 2 ml of CMF-PBS for 25-cm2 flask (for 75-cm2 flasks, use 5 ml CMF-PBS). Isolation and Characterization of Mesangioblasts

36. Replace rinse with 1 ml trypsin/EDTA for 25-cm2 flask (2 ml trypsin/EDTA for 75-cm2 flasks), and incubate 3 to 5 min at 37◦ C.

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37. Inactivate trypsin by adding 1 ml D20 medium in which all cells should be carefully aspirated. Pipet up and down two to three times to obtain a homogeneous cell suspension. 38. Centrifuge cells 5 min at 232 × g, room temperature. Discard supernatant. 39. Resuspend the pellet thoroughly in 6 to 8 ml of D20 medium and dispense 2-ml aliquots of cell suspension into each of three or four flasks (1:3 or 1:4 split, depending upon proliferation rate). Add D20 medium to reach a final volume of 5 ml for 25-cm2 flasks and 12 ml for 75-cm2 flasks. Drag the flasks with a cross-movement on the incubator shelf, to ensure homogeneous distribution of cells. Incubate at 37◦ C. When mesoangioblasts have been expanded to 25-cm2 flasks, they can be frozen for storage (see Support Protocol 4). In addition, detailed characterization of each clone should be performed at an early passage. For purpose of tracking passage number, begin counting passages the first time cells are plated without any feeder layer (i.e., step 34). Mouse embryonic aorta mesoangioblasts can be expanded up to 30 passages before showing signs of senescence.

ISOLATING, CLONING, PROPAGATING, AND FREEZING OF MOUSE ADULT MUSCLE MESOANGIOBLASTS

BASIC PROTOCOL 2

Murine adult mesoangioblasts differ from their embryonic counterpart in the expression of pericyte markers (such as alkaline phosphatase) and in the absence of endothelial markers (such as CD34). The slight differences in isolation and cloning of mesoangioblasts from mouse adult muscle, in comparison with embryonic aorta, are mainly due to the fact that primary cultures of adult tissues show a slower growth rate, and cloning efficiency may be lower. Mouse adult muscle fragments can be stored in D20 medium up to 24 hr at 4◦ C before being processed.

Materials Mouse skeletal (Tibialis anterior) muscle fragments (≥30 mg) Phosphate-buffered saline without Ca2+ /Mg2+ , (CMF-PBS; Sigma), sterile M5 medium (see recipe), sterile D20 medium (see recipe), sterile Collagenase/dispase solution (see recipe), sterile Trypan blue (Sigma) 0.05% (w/v) trypsin/0.02% (w/v) EDTA, (Sigma), sterile 6-, 10-, and 15-cm petri dishes (Nunc) Rounded-edge disposable scalpels, sterile Curved forceps, sterile 5% CO2 , 5% O2 , 90% N2 incubator Additional reagents and equipment for tissue processing (see Basic Protocol 1) Isolate and clone mouse adult muscle mesoangioblasts Follow the procedure described for mouse embryonic aorta (see Basic Protocol 1, steps 2 to 24) with the following step changes. 1. Dissect skeletal muscles of the mouse hind legs. 2. Rapidly rinse each skeletal muscle fragment in CMF-PBS to remove residual blood. 3. Perform dissections in a 10-cm petri dish containing 4 ml M5 medium. Dissect each fragment into 2-mm size pieces, trying to identify portions of interstitial tissue

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containing small vessels. Carefully remove adipose tissues, large nerves, and connective fascia. 4. Place five to ten pieces in collagen-coated 6-cm petri dish without feeder layer and 3 ml of D20 medium or scale up to larger dishes when processing larger amounts of tissue. 5. Incubate 5 to 8 days in a humidified chamber in a 37◦ C, 5% CO2 /5% O2 /90% N2 incubator. 6. Dissociate the cultures using 2 ml collagenase/dispase (see Basic Protocol 1, steps 13 to 19). Collect the cells in a 15-ml tube and centrifuge 15 min at 232 × g, room temperature. 7. Discard supernatant and resuspend the pellet in 300 µl of D20 medium. 8. Pipet up and down several times using a 1000-µl pipettor with filtered tips to disaggregate the muscle fragments as much as possible. Let the larger muscle debris sediment few seconds on the bottom of the tube and transfer the upper more homogeneous cell suspension to a new 15-ml centrifuge tube. 9. Count viable cells by trypan blue exclusion and proceed to cloning (see Basic Protocol 1, steps 22 to 26). 10. After 7 to 10 days, carefully inspect the cultures with a microscope to detect the first clones.

Propagate and freeze mouse adult muscle mesoangioblasts 11. Propagate (see Basic Protocol 1, steps 30 to 39) mesoangioblasts derived from mouse adult skeletal muscle and freeze (see Support Protocol 4) according to the same procedure described for mouse embryo aorta mesoangioblasts. ALTERNATE PROTOCOL 1

ISOLATION, PROPAGATION, AND CLONING OF HUMAN ADULT PERICYTE-DERIVED CELLS Human mesoangioblasts isolated from adult skeletal muscle have been more precisely defined as “pericyte-derived cells.” It has been observed that both the human and mouse adult counterpart of murine embryo mesoangioblasts express a series of pericyte markers, such as alkaline phosphatase and NG2, and do not express endothelial markers, such as CD34 (Dellavalle et al., 2007; Tonlorenzi, unpub. observ.). It is likely that adult pericytederived cells originate from embryonic mesoangioblasts but this has not been formally demonstrated.

Additional Materials (also see Basic Protocol 1) Skeletal muscle fragments (≥100 mg) from a muscle biopsy M5 medium (see recipe), sterile Prepare skeletal muscle fragment 1. Rapidly rinse each skeletal muscle fragment in CMF-PBS to remove residual blood. 2. Dissect the muscle in a 10-cm petri dish containing 4 ml M5 medium. Dissect each fragment into 2-mm pieces, trying to identify portions of interstitial tissue containing small vessels. Remove fat where present. No care is taken to clean vessels from surrounding mesenchyme and segments of muscle fibers. Isolation and Characterization of Mesangioblasts

It is important to remove as much adipose tissue as possible, since its presence may delay cell outgrowth.

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3. Pretreat the appropriate number of 6-cm collagen-coated petri dishes by pipetting 3 ml M5 medium into each dish, making sure that the surface is completely covered. Gently aspirate the medium gently, but not completely, so that the dish surface remains thoroughly wet. 4. Transfer the selected fragments (four to five fragments) into each 6-cm dish in a drop of medium that is created between the arms of a curved forceps. Scale up to 10-cm collagen-coated petri dishes if processing larger amounts of tissue.

5. After the fragments have been transferred into the dishes, add 2 ml M5 medium (3.5 ml for 10-cm petri dish) pipetting it along the edge of the dish to prevent detachment and floating of fragments.

Culture cells 6. Prepare a humidified chamber by placing the dishes into a clean plastic box, along with two open 6-cm petri dishes filled with sterile distilled water. Cover the box with aluminum foil. Incubate overnight in a 37◦ C, 5% CO2 /5% O2 /90% N2 humidified incubator. 7. Around 24 hr after initiation of cultures, carefully add an additional 2 ml M5 medium to each dish (3 to 4 ml for 10-cm petri dishes). A dedicated incubator should be used, or at least a rarely opened incubator.

8. After 5 to 7 days, examine the cultures for preliminary growth of adherent cells. 9. Add 1 to 2 ml of freshly prepared, prewarmed M5 medium to each dish (a 6-cm petri dish easily contains 6 ml). 10. After an additional 2 to 3 days, examine the cultures for pericyte-derived cells, which are distinguishable as small, round, very refractile cells, floating or weakly adhering to the layer of flat cells below. 11. Carefully transfer culture medium and floating cells to a new, uncoated petri dish of the same size as used for primary culture. Add freshly prepared, pre-warmed medium to reach a total volume of 5 ml for 6-cm petri dish (10 ml for 10-cm petri dish). Gentle pipetting may help to detach the weakly adhering cells around the explants. In case of poor recovery of floating cells, transfer to a smaller petri dish (see Troubleshooting).

12. After 24 hr, examine the cultures to see that ∼50% to 70% of the floating cells adhere to the plastic surface. A floating fraction should always be clearly distinguishable.

Trypsinize cells 13. When the adherent fraction of the cell population reaches 70% to 80% confluence, proceed to trypsinization and transfer to flasks. Due to variability in cell proliferation rate, 70% to 80% confluence of adherent fraction may require 2 to 4 days. The culture does not need to be fed between trypsinizations.

Propagate human pericyte-derived cells 14. At 70% to 80% confluence of the adherent cell population, remove culture medium and set aside in 15-ml centrifuge tubes. Floating and adherent cells do not differ since floating cells, cultured separately, will give rise to ∼50% adherent cells and vice versa.

15. Rinse the growing surface with 2 ml CMF-PBS.

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16. Add 2 ml of trypsin/EDTA and incubate 3 to 5 min at room temperature. Check under a microscope for complete detachment of cells. Human pericyte-derived cells are very sensitive to trypsin. If cells are healthy, their detachment should be very quick and complete.

17. Use the medium set aside (step 14) to collect all cells. In this way, both floating and adherent populations are simultaneously recovered.

18. Centrifuge 10 min at 232 × g, room temperature. 19. Gently resuspend the pellet in 6 ml M5 medium and dispense 2-ml aliquots of cell suspension into each of three flasks (1:3 split). Add M5 medium to reach a final volume of 5 ml for 25-cm2 tissue culture flasks and 12 ml for 75-cm2 tissue culture flasks. The day after trypsinization, the floating population may be reduced. Normally, this fraction should start to expand again after 48 hr. When pericyte-derived cells have been expanded to 75-cm2 tissue culture flasks, proceed to characterization and karyotype analysis. In addition, it is recommended to freeze (see Support Protocol 4) several vials of cells at early passages for future use and further propagation. To keep a record of passage number, begin counting passages at first trypsinization. Human pericyte-derived cells can be expanded up to 20 passages under 5% O2 tension. At pre-senescence a strong reduction in the floating population of cells is observed, in addition to the presence of large, flat or elongated, vacuolated cells.

Clone human pericyte-derived cells Human pericyte-derived cells, derived and grown under physiological O2 tension, can be cloned at very early passages (2 to 4 passages) without the support of any feeder layer. Cells selected for cloning have to be detached during the proliferating phase (∼48 hr after trypsinization). The expected cloning efficiency is usually 1% to 2%. Culture medium used for cloning experiments must be freshly prepared. 20. Remove culture medium from one 25-cm2 flask of human pericyte-derived cells and set it aside. 21. Rinse the growing surface with 2 ml CMF-PBS. 22. Add 2 ml trypsin/EDTA. Incubate 3 to 5 min at room temperature. 23. Collect detached cells and add to saved medium. Centrifuge 5 min at 242 × g, room temperature. 24. Suspend the pellet in 2 ml M5 medium, and count viable cells by trypan blue exclusion, using a hemacytometer (UNIT 1C.3). 25. Dilute cells in M5 medium to obtain 150 ml of each of the following concentrations:

1 cell/ml 5 cells/ml 10 cells/ml A total volume of ∼150 ml of cell suspension is enough for three 48-well plates.

Isolation and Characterization of Mesangioblasts

26. For each concentration, plate cell suspensions at 1 ml/well in three 48-well plates. As a control of cell proliferation, plate 100 cells/well in a few wells of a separate 48-well plate.

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27. Prepare a humidified chamber by placing the 48-well plates into a clean plastic box along with two open 6-cm petri dishes filled with sterile distilled water. Cover the box with aluminum foil. 28. Place for at least 1 week in a 37◦ C, 5% CO2 /5% O2 /90% N2 humidified incubator. A dedicated incubator should be used, or at least a rarely opened incubator.

29. After 8 to 10 days, examine the plates carefully under a microscope to detect the first clones. Longer incubations (up to 15 days) may be necessary.

30. Add 200 µl of fresh medium to each well on day 7 or 8. 31. Passage the clones using trypsin/EDTA when the cells have covered at least 50% of the well surface. After the first trypsinization, do not split the cells, but plate them in a new well of the same size. From second passage on, proceed to 1:2 splitting.

32. When clones have been expanded to 25-cm2 tissue culture flasks, proceed to differentiation tests and karyotype analysis. Clones of human cells normally have a reduced lifespan. Typical pre-senescent cells (Fig. 2B.1.1G) may appear after passage 13 to 15.

ISOLATION AND CLONING OF ADULT DOG SKELETAL MUSCLE MESOANGIOBLATS

ALTERNATE PROTOCOL 2

Adult dog skeletal muscle mesoangioblasts can be isolated, propagated, and cloned according to the same procedures described for human skeletal muscle. Morphologically, established cultures of dog mesoangioblasts are characterized by a smaller fraction of floating cells with respect to the corresponding human pericyte-derived cells during the proliferation phase. Because of the higher proliferation rate of dog in comparison with human cells, incubation times for tissue explants may be shorter (3 to 7 days may be sufficient for mesoangioblast outgrowth). As far as cell propagation is concerned, dog postnatal skeletal muscle mesoangioblasts can be expanded for ∼25 passages before senescence and, differently from human cells, can be split at a higher ratio (up to 1:5). As with human pericyte-derived cells, canine mesoangioblast clones have a reduced lifespan (∼10 to 12 passages), which can be slightly extended by culture under physiological O2 tension (up to 15 passages).

DIFFERENTIATION OF MESOANGIOBLASTS: CO-CULTURE WITH MURINE C2C12 MYOBLASTS

BASIC PROTOCOL 3

This assay tests the ability of mesoangioblasts to differentiate into skeletal muscle cells (Minasi et al., 2002) in the presence of an inducer cell line, such as C2C12 (mouse myoblasts) or L6 (rat myoblasts; see Alternate Protocol 3). To be easily distinguished, the mouse mesoangioblasts to be tested in co-cultures should be previously transduced with a lentiviral vector expressing nuclear-LacZ; in the case of rat myoblasts, mouse and rat nuclei can be distinguished by DAPI staining that reveals speckles in the mouse nucleus but not in the rat. The co-culture assay is best performed using C2C12 cells with murine mesoangioblasts, and L6 cells with canine mesoangioblasts or human pericyte-derived cells. L6 cells, in contrast to C2C12, can withstand prolonged culture in differentiation conditions that are usually necessary for human or dog cell differentiation. C2C12-derived myotubes tend to detach from the dish after several days.

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C2C12 and L6 cells to be used in co-culture experiments should be in good condition and proliferating well (see Troubleshooting). This protocol is used to induce skeletal myogenic differentiation in murine mesoangioblasts by co-culturing them with C2C12 murine myoblasts.

Materials C2C12 cells grown in a 25-cm2 tissue culture flask (ATCC #CRL-1772) Phosphate-buffered saline without Ca2+ /Mg2+ (CMF-PBS; Sigma), sterile 0.05% (w/v) trypsin/0.02% (w/v) EDTA (Sigma), sterile D20 medium (see recipe) D10 medium (see recipe) Mesoangioblasts to be tested D2 medium (see recipe) 4% (w/v) paraformaldehyde (PFA) 37◦ C, 5% CO2 incubator 3.5-cm petri dishes Additional reagents and equipment for trypan blue exclusion (UNIT 1C.3) Prepare C2C12 dishes 1. Aspirate and discard medium from C2C12 flask, and rinse the cells with 2 ml CMFPBS. 2. Replace with 1 ml of trypsin/EDTA and incubate 3 to 5 min in a 37◦ C, 5% CO2 incubator. 3. Inactivate trypsin by adding 3 ml D20 medium down the growing surface to collect all cells. Pipet up and down two to three times to obtain a homogeneous solution. To obtain uniformly dispersed C2C12 cells, trypsinization has to be complete. No clumps should appear in the cell suspension.

4. Centrifuge 5 min at 232 × g, room temperature, to pellet cells. Discard supernatant. Carefully resuspend pellet in 5 ml D20 medium. Count viable cells by trypan blue exclusion, using a hemacytometer (UNIT 1C.3). 5. Plate 5 × 104 cells /3.5-cm petri dish in 2 ml D10 medium. For each mesoangioblast line to be tested, at least three dishes have to be plated with C2C12 cells.

6. Incubate 2 hr to overnight in a 37◦ C, 5% CO2 incubator.

Prepare mesoangioblasts 7. Detach and count mesoangioblasts to be tested, following the procedure described for C2C12 (steps 1 to 4). 8. Dilute mesoangioblasts to 104 cells/ml in D20 medium.

Initiate co-cultures 9. Remove medium from C2C12 dishes.

Isolation and Characterization of Mesangioblasts

10. Immediately plate 0.5 ml and 1 ml of mesoangioblasts suspension onto first and second C2C12 dish to obtain a mesoangioblast/C2C12 ratio of 1:10 and 1:5, respectively. Bring volume up to 2 ml with D20 medium. Replace D10 medium of the third C2C12 dish with 2 ml D20 medium without any addition of mesoangioblasts (control dish).

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11. Incubate overnight in a 37◦ C, 5% CO2 incubator. Drag the dishes with a cross-movement on the incubator shelf, to ensure homogeneous distribution of mesoangioblasts over the C2C12 layer.

12. On the following day, replace D20 medium with 2 ml D2 medium (low-serum differentiation medium) in all of the three C2C12 dishes. 13. Monitor differentiation on C2C12 control dish beginning on day 3. Myotubes should start to be evident after 3 to 4 days of incubation as long, multinucleated cells (see Fig. 2B.1.1H). Differentiation should be complete after 5 to 7 days on co-culture dishes.

14. Remove medium from petri dishes, and carefully rinse the growing surface with 1 ml of CMF-PBS. Gentle aspiration is recommended, since the very confluent growing surface may easily detach as a whole layer.

15. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove PFA. Rinse with 1 ml CMF-PBS. Proceed immediately to immunofluorescence, or store up to 48 hr at 4◦ C. If dishes need to be stored, add 0.5 ml PBS to each dish and seal with Parafilm to avoid liquid evaporation and/or contamination.

16. Calculate the percentage of myogenic differentiation as the number of mesoangioblast nuclei [detected by DAPI (Lin et al., 1976) or X-gal (Cepko, 1996) staining] inside myosin-positive cells or myotubes [immunostained (Minasi et al., 2002) with MF20 monoclonal antibody, from Developmental Hybridoma Bank, that recognizes all sarcomeric myosin heavy chains] divided by the total number of mesoangioblast nuclei and multiplied by 100. In the case of n-LacZ, data may be overestimated since the nuclear LacZ synthesized in the cytoplasm may be targeted to a neighbor C2C12 nucleus. This is not the case for DAPI staining.

DIFFERENTIATION OF CANINE MESOANGIOBLASTS OR HUMAN PERICYTE-DERIVED CELLS: CO-CULTURE WITH L6 RAT MYOBLASTS

ALTERNATE PROTOCOL 3

Canine mesoangioblasts and human pericyte-derived cells are co-cultured with rat L6 myoblasts to induce differentiation. Human and dog cells can be efficiently detected by the use of a specific antibody, directed against human nuclear lamin A/C (Novocastra, cat. no. NCL-LAM A/C), that cross-reacts with dog, but not with rodent cells.

Additional Materials (also see Basic Protocol 3) L6 cells grown in a 25-cm2 tissue culture flask Mesoangioblasts to be tested grown in a 25-cm2 tissue culture flask M5 medium (see recipe) 15-ml centrifuge tubes Prepare L6 dishes 1. Aspirate and discard medium from L6 flask, and rinse with 2 ml CMF-PBS. 2. Replace with 1 ml trypsin/EDTA and incubate 3 to 5 min in a 37◦ C, CO2 incubator. 3. Inactivate trypsin by adding 3 ml D20 medium down the growing surface to collect all cells. Pipet up and down two to three times to obtain a homogeneous solution. Somatic Stem Cells

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4. Centrifuge 5 min at 242 × g, room temperature, to pellet. Discard supernatant. Carefully resuspend pellet in 5 ml D20 medium. Count viable cells by trypan blue exclusion, using a hemacytometer (UNIT 1C.3). 5. Plate 5 × 104 cells /3.5-cm petri dish in 2 ml D10 medium. For each mesoangioblast line to be tested, at least three dishes have to be plated with L6 cells.

6. Incubate 2 hr or overnight in a 37◦ C, CO2 incubator.

Prepare mesoangioblasts/pericyte-derived cells 7. Remove medium from mesoangioblasts/pericyte-derived cells and set it aside in a 15-ml centrifuge tube. 8. Rinse the growing surface with 2 ml CMF-PBS. Add 2 ml trypsin/EDTA and incubate 3 to 5 min at room temperature. 9. Collect the cells and add to the medium saved in a 15-ml centrifuge tube from step 7. 10. Centrifuge 10 min at 232 × g, room temperature. 11. Gently resuspend the pellet in 5 ml M5 medium and count viable cells by trypan blue exclusion using a hemacytometer (UNIT 1C.3). 12. Dilute cells to 104 /ml in M5 medium.

Initiate co-cultures 13. Remove medium from L6 dishes. 14a. For canine mesoangioblasts: Immediately plate 0.5 ml and 1 ml of canine mesoangioblast suspension onto first and second L6 dishes, to obtain a mesoangioblasts/L6 ratio of 1:10 and 1:5, respectively. 14b. For human pericyte-derived cells: In the case of human pericyte-derived cells, plate 1 ml and 1.5 ml of cell suspension onto the first and second L6 dish to obtain ratios of 1:5 and ∼1:3 with L6 cells, respectively. 15. Bring volume up to 2 ml with M5 medium. Replace D10 medium of the third L6 dish with 2 ml M5 medium without any addition of mesoangioblasts or pericyte-derived cells (control dish). 16. Incubate 24 hr in a 37◦ C, CO2 incubator. Drag the dishes with a cross-movement on the incubator shelf to ensure homogeneous distribution of the mesoangioblast/pericyte-derived cells over the L6 layer.

17. On the following day, remove M5 medium from each dish and wash the growing surface two times with 1 ml of CMF-PBS, each time. 18. Add 2 ml D2 medium (low-serum differentiation medium) in all of the three L6 dishes.

Monitor differentiation 19. Monitor differentiation on L6 control dish beginning on day 3. Myotubes should start to be evident after 3 to 4 days of incubation. Differentiation on co-culture dishes should be complete after 6 to 8 days (dog mesoangioblasts) or 7 to 10 days (human pericyte-derived cells). Isolation and Characterization of Mesangioblasts

For incubation times longer than 7 days, add 0.5 ml fresh D2 medium to each co-culture dish. For human pericyte-derived cell co-cultures, incubation time may be extended to 12 days.

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20. Remove medium from petri dishes, and carefully rinse the growing surface with 1 ml CMF-PBS. Mild aspiration is recommended, since the very confluent growing surface may easily detach as a whole layer.

21. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove 4% PFA and rinse with 1 ml CMF-PBS. Proceed immediately to immunofluorescence, or store up to 48 hr at 4◦ C. If dishes are to be stored, add 0.5 ml PBS to each dish and seal with Parafilm to avoid drying out and/or contamination.

22. Calculate percentage of myogenic differentiation as the number of mesoangioblast/pericyte-derived cell nuclei (detected by staining with anti-lamin A/C antibody) inside myosin-positive cells or myotubes (stained with MF20 antibody) divided by the total number of mesoangioblast/pericyte-derived cell nuclei multiplied by 100 (Fig. 2B.1.1M,Q).

DIFFERENTIATION OF HUMAN PERICYTE-DERIVED CELLS: SPONTANEOUS SKELETAL MYOGENIC DIFFERENTIATION

ALTERNATE PROTOCOL 4

In contrast to murine mesoangioblasts, human pericyte-derived cells and, to a lower extent, dog mesoangioblasts can spontaneously differentiate into multinucleated skeletal myotubes (Dellavalle et al., 2007) when cultured onto a Matrigel-coated plastic support (see Support Protocol 3).

Additional Materials (also see Basic Protocol 3) Dog mesoangioblasts/human pericyte-derived cells to be tested grown in a 25-cm2 tissue culture flask M5 medium (see recipe) Reduced growth factor Matrigel–coated 3.5-cm petri dishes (see Support Protocol 3), freshly prepared 1. Detach and count dog mesoangioblasts/human pericyte-derived cells according to Alternate Protocol 3, steps 7 to 12. 2. Plate 5 × 104 cells/Matrigel-coated 3.5-cm petri dish in 2 ml M5 medium. Due to variability in cell proliferation rate and differentiation efficiency, slight adjustment in cell number/dish may be necessary (5 × 104 –105 cells/dish).

3. Incubate overnight in a 37◦ C, 5% CO2 incubator. 4. Remove medium and rinse each dish with 1 ml CMF-PBS. 5. Add 2 ml D2 differentiation medium to each dish. 6. Incubate at least 1 week in a 37◦ C, 5% CO2 incubator. At that time, first myotubes should be evident on test dishes (for both canine and human cells). A time period of 7 to 8 days are usually sufficient for canine mesoangioblast differentiation, while 10 to 12 days may be necessary for human pericyte-derived cells (Fig. 2B.1.1H).

7. Remove medium from petri dishes, and carefully rinse the growing surface with 1 ml CMF-PBS. Mild aspiration is recommended, since the very confluent growing surface may easily detach as a whole layer.

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8. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove 4% PFA. Rinse with 1 ml CMF-PBS. Proceed immediately to immunofluorescence, or store up to 48 hr at 4◦ C. If dishes need to be stored, add 0.5 ml PBS to each dish and seal with Parafilm to avoid drying out and/or contamination.

9. Calculate percentage of myogenic differentiation as the number of mesoangioblast/pericyte-derived cell nuclei (detected by DAPI) inside myosinpositive cells or myotubes divided by the total mesoangioblast nuclei multiplied by 100. ALTERNATE PROTOCOL 5

DIFFERENTIATION OF MESOANGIOBLAST/PERICYTE-DERIVED CELLS: INDUCTION OF SMOOTH MUSCLE CELL DIFFERENTIATION BY TGFβ TREATMENT Murine and canine mesoangioblasts and human pericyte-derived cells are very sensitive to TGFβ treatment, which is known to induce smooth muscle differentiation (Ross et al., 2007). The suggested final concentration to be adopted in differentiation medium is 5 ng/ml. Suggested differentiation time is ∼6 to 7 days even for human cells. Morphological change to large, flat and typically elongated cells should be evident starting from day 3 to day 4.

Additional Materials (also see Basic Protocol 3) Mesoangioblasts/human pericyte-derived cells to be tested grown in a 25-cm2 tissue culture flask M5 medium (see recipe) 5 µg/ml TGFβ stock solution Prepare test cultures 1. Detach and count mesoangioblasts/human pericyte-derived cells according to Basic Protocol 3, steps 7 to 8 for murine mesoangioblasts or Alternate Protocol 3, steps 7 to 12 for dog mesoangioblasts and human pericyte-derived cells. 2. Plate 5 × 104 cells/3.5-cm petri dish in 2 ml of corresponding medium (D20 medium for murine, M5 medium for canine and human cells). For each cell line to be tested, plate at least two dishes (one test and one control dish). 3. Incubate overnight in a 37◦ C, 5% CO2 incubator.

Differentiate cultures 4. Remove medium and rinse each dish with 1 ml CMF-PBS. 5. Add 1.5 ml D2 differentiation medium to each dish. 6. Add 1.5 µl TGFβ stock solution to each test dish (5 ng/ml final). 7. Add 1.5 µl fresh TGFβ stock solution every other day. TGFβ concentrated stock solution (5 µg/ml) can be stored up to 10 days at 4◦ C. For longer storage, small aliquots should be frozen at −20◦ C. Once thawed, TGFβ solution cannot be frozen again.

8. Check cultures for smooth muscle differentiation, which should be complete after 6 to 7 days (six or seven total additions of TGFβ). Isolation and Characterization of Mesangioblasts

Fix and score the cells for smooth muscle differentiation 9. Remove medium from petri dishes and carefully rinse the growing surface with 1 ml CMF-PBS.

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10. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove 4% PFA. Rinse with 1 ml CMF-PBS. Proceed immediately to immunofluorescence, or store up to 48 hr at 4◦ C. If dishes need to be stored, add 0.5 ml PBS to each dish and seal with Parafilm to avoid drying out and/or contamination.

11. Calculate percentage of smooth muscle differentiation as the number of mesoangioblast cells expressing a smooth muscle phenotype (detected by an antibody directed against smooth alpha actin (Sigma cat. no. A2547) or calponin (Sigma cat. no. C2687) divided by total number of mesoangioblast nuclei multiplied by 100 (Fig. 2B.1.1J).

DIFFERENTIATION OF MESOANGIOBLAST/PERICYTE-DERIVED CELLS: INDUCTION OF OSTEOBLAST DIFFERENTIATION BY BMP2 TREATMENT

ALTERNATE PROTOCOL 6

The effect of BMP2 treatment is particularly evident on mouse embryo aorta mesoangioblasts, since these cells do not normally express alkaline phosphatase (Fig. 2B.1.1K,O), while adult dog mesoangioblasts and human pericyte-derived cells usually do (Fig. 2B.1.1L). Nevertheless, even on canine and human cells, BMP2 treatment results in further increase of alkaline phosphatase activity (Fig. 2B.1.1P).

Additional Materials (also see Basic Protocol 3) Mesoangioblast/human pericyte-derived cell cultures to be tested grown in a 25-cm2 tissue culture flask 10 µg/ml BMP2 stock solution Alkaline phosphatase staining solution (see recipe), freshly prepared Establish test cultures 1. Detach and count mesoangioblast/human pericyte-derived cells according to Basic Protocol 3, steps 7 to 8, for murine mesoangioblasts or Alternate Protocol 3, steps 7 to 12, for dog mesoangioblasts and human pericyte-derived cells. 2. Plate 5 × 104 cells/3.5-cm petri dish in 2 ml of appropriate medium (D20 medium for murine cells, M5 medium for canine and human cells). For each cell line to be tested, plate at least two dishes (one test and one control dish). 3. Incubate overnight in a 37◦ C, 5% CO2 incubator.

Differentiate cultures 4. Remove medium and rinse each dish with 1 ml CMF-PBS. 5. Add 1.5 ml D2 differentiation medium to each dish. 6. Add 15 µl BMP2 stock solution to each test dish (100 ng/ml final concentration). No addition has to be made to control dishes.

7. On every other day, add 15 µl fresh BMP2 stock solution to test dishes. BMP2 concentrated stock solution (10 µg/ml) can be stored up to 10 days at 4◦ C. For longer storage, small aliquots should be frozen at −20◦ C. Once thawed, BMP2 solution cannot be frozen again.

8. Assess the cultures for differentiation, which should be complete after 6 to 7 days (six or seven total additions of BMP2). Somatic Stem Cells

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Fix and score cultures for osteoblast differentiation 9. Remove medium from petri dishes, and carefully rinse the growing surface with 1 ml CMF-PBS. 10. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove 4% PFA. Rinse with 1 ml CMF-PBS. Proceed immediately to alkaline phosphatase staining. Fixed cultures may be stored up to 48 hr at 4◦ C. If stored, add 0.5 ml CMF-PBS to each dish and seal with Parafilm to avoid drying out and/or contamination.

11. Remove CMF-PBS and add 1 ml of alkaline phosphatase staining solution to each test and control dish. 12. Incubate 2 hr at room temperature in the dark. 13. Examine cultures under inverted phase-contrast microscope for a brown cytoplasmic stain, whose intensity is roughly proportional to the level of enzymatic activity. For a more rigorous test of osteoblast differentiation, in vitro formation of Von Kossa positive, calcified nodules (Chaplin and Grace, 1975) should be characterized. ALTERNATE PROTOCOL 7

DIFFERENTIATION OF MESOANGIOBLAST/PERICYTE-DERIVED CELLS: INDUCTION OF ADIPOCYTE DIFFERENTIATION The use of adipogenic induction medium permits a test of the mesoangioblast/pericytederived cell potential to give rise to adipose cells. Oil Red O is a lysochrome fat-soluble dye used for staining of neutral triglycerides. If cells grown in this medium differentiate into adipocytes, their triglyceride content is stained intensely red by the Oil Red O treatment.

Additional Materials (also see Basic Protocol 3) Mesoangioblast/human pericyte-derived cell cultures to be tested grown in a 25-cm2 tissue culture flask Adypogenic induction medium (Cambrex) Oil Red O solution (see recipe) Inverted phase-contrast microscope Prepare cultures 1. Detach and count mesoangioblast/human pericyte-derived cells according to Basic Protocol 3, steps 7 to 8, for murine mesoangioblasts or Alternate Protocol 3, steps 7 to 12, for dog mesoangioblasts and human pericyte-derived cells. 2. Plate 5 × 104 cells/3.5-cm petri dish in 2 ml of corresponding medium (D20 medium for murine, M5 medium for canine and human cells). For each cell line to be tested, plate at least two dishes (one test and one control dish). 3. Incubate overnight in a 37◦ C, 5% CO2 incubator.

Add adipogenic medium 4. Remove medium and rinse each dish with 1 ml CMF-PBS. 5. Add 1.5 ml of adypogenic induction medium to each test dish. Add 1.5 ml D2 medium to each control dish. 6. Check cultures for differentiation after 6 to 7 days for murine and canine mesoangioblasts or up to 10 days for human pericyte-derived cells. Isolation and Characterization of Mesangioblasts

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Differentiation to adipocytes is morphologically easy to detect. It is characterized by the presence of gradually enlarging, translucent vacuoles in the cytoplasm of a percentage of cells (up to 60% to 70%). The presence of lipid content in these vacuoles must be confirmed by appropriate staining (Oil-Red O staining, as shown in Fig. 2B.1.1N). Current Protocols in Stem Cell Biology

7. Remove medium from petri dishes, and carefully rinse the growing surface with 1 ml CMF-PBS.

Stain cells with Oil Red O 8. Fix with 1 ml of 4% PFA 5 min at room temperature. Remove 4% PFA. Rinse with 1 ml CMF-PBS. Proceed immediately to Oil-Red O staining. 9. Remove CMF-PBS and add 1 ml Oil-Red O solution to each test and control dish. 10. Incubate 2 hr at room temperature. 11. Remove and discard Oil-Red O solution. 12. Carefully rinse the culture surfaces two to three times with 1 ml distilled water.

Score cells for adipocyte differentiation 13. Finally, add 500 µl distilled water to prevent surface from drying out, and proceed to analyze the dishes under an inverted phase-contrast microscope. Cells containing one or more brightly stained vesicles are counted as differentiated adipocytes.

PREPARATION OF MITOTICALLY INACTIVE STO FEEDER LAYER Many mammalian cells proliferate poorly under clonal conditions; their growth is significantly enhanced by supporting cells (feeder), whose mitotic activity has been previously arrested by chemical (mytomicin C) or physical (X-ray) means. STO cells are an established mouse embryo fibroblast cell line commonly used as a feeder layer.

SUPPORT PROTOCOL 1

CAUTION: MMC is light sensitive and highly toxic. This substance must be handled with care in the dark; refer to the manufacturer’s product datasheet for instructions.

Materials Frozen vials of STO (ATCC # CRL-1503) D10 medium (see recipe) 0.05% (w/v) trypsin/0.02% (w/v) EDTA (Sigma), sterile Mitomycin C stock solution (MMC, see recipe) Phosphate-buffered saline without Ca2+ /Mg2+ (CMF-PBS; Sigma), sterile 15-ml centrifuge tube 75-cm2 vented tissue culture flasks (Nunc) 37◦ C, 5% CO2 incubator Additional reagents and equipment for trypan blue staining (UNIT 1C.3) Thaw STO cells 1. Rapidly thaw a frozen vial of STO cells at 37◦ C. 2. Transfer the cells to a 15-ml centrifuge tube containing 5 ml D10 medium and centrifuge 5 min at 232 × g, room temperature. 3. Resuspend the cell pellet in 12 ml D10 medium and plate in a 75-cm2 vented tissue culture flask. Incubate overnight in a 37◦ C, 5% CO2 incubator.

Mitotically inactivate cells 4. On the following day, split the cells 1:5 using trypsin/EDTA treatment as described for C2C12 (see Basic Protocol 3). Somatic Stem Cells

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At 60% to 70 % confluence, proceed to cell inactivation by mitomycin C (MMC) treatment (step 5). Cell density is crucial to perform an efficient inactivation. If cells reach a higher density, a new splitting is necessary before MMC treatment. CAUTION: MMC is light sensitive and highly toxic. Handle with care in the dark.

5. Remove medium from each flask and add 10 ml D10 medium containing 100 µl of MMC (1 mg/ml), making sure that the entire growing surface is covered. 6. Incubate cells 3 hr in a 37◦ C, 5% CO2 incubator. 7. Remove the medium and carefully rinse the cell monolayer with 10 ml CMF-PBS three times.

Prepare feeder plates 8. Add 2 ml of trypsin/EDTA to each flask and incubate 5 to 10 min at 37◦ C, monitoring the complete detachment of cells under a microscope. 9. Add 8 ml D10 medium to each flask and carefully collect all cells. Gently pipet up and down two to three times to disaggregate any remaining cell clumps. The high volume of medium is necessary to perform a further rinse of the MMC-treated cells.

10. Centrifuge cells 10 min at 242 × g, room temperature. 11. Resuspend each cell pellet in 2 ml D10 medium and count viable cells using a trypan blue hemacytometer (UNIT 1C.3). 12. Immediately plate the cells at 1–1.5 × 104 cells/cm2 (∼1–1.5 × 104 cells/well for a 48-well plate). Plate cells in a volume of 1 ml/well in a 48-well plate.

13. Allow the cells to attach 6 to 8 hr or overnight in a 37◦ C, 5% CO2 incubator. Use the MMC-inactivated cells within 36 hr. Each plate should be checked under a microscope before being used in cloning experiments. If cells are viable and have been counted properly, they should appear as a subconfluent layer in each well. Some strains of cells currently used as feeder layers may exhibit different sensitivity to MMC inactivation. A titration should be performed to determine the effective MMC dose, performing the inactivation both in presence and absence of FBS in culture medium. SUPPORT PROTOCOL 2

COLLAGEN COATING OF TISSUE CULTURE SURFACES It is essential that, at the moment of use, collagen-coated petri dishes are completely dry. Therefore, it is recommended to prepare them 24 hr in advance. Once dry, collagen-coated petri dishes can be stored up to 3 months at 30◦ C. CAUTION: Collagen solution used for coating contains 20% (v/v) acetic acid and should be used in a chemical hood.

Materials Isolation and Characterization of Mesangioblasts

Collagen type I solution (see recipe) Petri dishes 30◦ C oven

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1. Place the appropriate number of petri dishes to be coated in a chemical hood. 2. Carefully add the collagen type I solution into each petri dish making sure the whole surface is completely covered. Use 1 ml, 5 ml, and 10 ml of collagen solution for 3.5-, 6-, and 9-cm petri dish, respectively.

3. Let stand 5 min. 4. Slowly remove ∼80% to 90% of the solution, leaving the surface of the dish uniformly wet. Collagen solution can be recycled and used several times; therefore, it has to be carefully removed with a pipet and not by vacuum aspiration.

5. Transfer collagen-coated petri dishes to dedicated 30◦ C oven. Once completely dry, collagen-coated dishes can be used or can be stored up to 3 months in the 30◦ C oven.

MATRIGEL COATING OF TISSUE CULTURE SURFACES Matrigel-coated petri dishes have to be freshly prepared and cannot be stored. Matrigel supports myogenic differentiation better than collagen but is more expensive.

SUPPORT PROTOCOL 3

Materials Reduced growth factors Matrigel stock solution (BD Biosciences; see recipe) High-glucose DMEM (Sigma), ice cold Petri dishes 37◦ C, 5% CO2 incubator 1. Thaw Matrigel stock solution on ice. 2. Prepare the working solution by diluting the stock 1:80 in cold DMEM (without any supplement). Matrigel working solution can be stored up to 24 hr at 4◦ C.

3. Place the appropriate number of petri dishes to be coated under a hood. 4. Apply Matrigel working solution carefully into each petri dish making sure the whole surface is completely covered. Use 1 ml, 3 ml, and 7 ml of Matrigel working solution for 3.5-, 5-, and 9-cm petri dishes, respectively.

5. Incubate 30 min in a 37◦ C, 5% CO2 incubator. 6. Remove and discard the Matrigel working solution, leaving the dish surface slightly wet. 7. Rinse the dish surface with appropriate culture medium before plating cells. 8. Use the Matrigel-coated petri dishes immediately.

FREEZING MESOANGIOBLASTS AND HUMAN PERICYTE-DERIVED CELLS Mesoangioblasts and human pericyte-derived cells proliferate for a limited number of passages; it is thus necessary to freeze cells from early passages to maintain a stock of the cell line. Murine, canine mesoangioblasts and human pericyte-derived cells are frozen following the same procedure.

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Materials Murine and canine mesoangioblasts cultures grown in a 25-cm2 tissue culture flask or human pericyte-derived cell cultures grown in a 75-cm2 tissue culture flask D20 or M5 medium (see recipes) Freezing solution (see recipe), freshly prepared, ice cold 0.05% (w/v) trypsin/0.02% (w/v) EDTA (Sigma), sterile DMSO (Sigma), sterile Hemacytometer 1.8-ml sterile cryovials (Corning), ice cold Cryogenic-controlled rate freezing container (Nalgene) or insulated cardboard/polystyrene foam box 1. Detach cells with trypsin/EDTA according to corresponding steps described for cell propagation (see Basic Protocols 1 and 2). Cells should be healthy and at 70% to 80% confluent at time of freezing. Mitosis should be evident under microscope inspection.

2. Suspend the cell pellet in 5 ml of appropriate medium (D20 medium for murine mesoangioblasts and M5 medium for dog mesoangioblasts and human pericytederived cells). 3. Count cells with a hemacytometer (UNIT 1C.3). 4. Centrifuge 5 min at 232 × g, room temperature. 5. Discard supernatant and gently suspend cells in appropriate volume of cold freezing solution to obtain the following cell concentration:

1–3 × 106 cells/mlfor murine mesoangioblasts 1–2 × 106 cells/mlfor canine mesoangioblasts and human pericyte-derived cells. 6. Set up the appropriate number of 1.8-ml cryovials and dispense 1 ml of cell suspension into each. Each cryovial should be clearly labeled with date, cell line code, and passage number.

7. Transfer vials into a freezing container and place overnight at −80◦ C. 8. On the following day, transfer vials to −135◦ C or to a liquid nitrogen container. Upon thawing, which is performed quickly in a 37◦ C water bath, transfer the vial content into a 15-ml centrifuge tube containing 5 ml of prewarmed appropriate culture medium, centrifuge 5 min at 232 × g, room temperature, discard supernatant to remove DMSO, resuspend cells in appropriate medium prior to plating of cells.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. NOTE: Fetal bovine serum (FBS, Cambrex, cat. no. DE14-801F) and horse serum (HS, Euroclone, cat. no. ECS0090L) used for media supplementation and in protocol steps have to be heatinactivated for 45 min at 56◦ C prior to use.

Alkaline phosphatase staining solution Isolation and Characterization of Mesangioblasts

4.5 µl/ml 4-nitroblue tetrazolium chloride (3 mg/ml NTC, Roche cat. no. 11383213001) continued

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3.5 µl/ml 5-bromo-4-chloro-3-indolyl-phosphate (50 mg/ml BCIP, Roche cat. no. 1383221) Buffered solution, pH 9.5 (see recipe) Prepare fresh Particular attention must be paid to the exact pH (9.5) of the buffered solution.

Buffered solution, pH 9.5 100 mM NaCl 100 mM Tris·Cl, pH 9.5 50 mM MgCl2 0.1% (v/v) Tween 20 Adjust pH with 0.1N HCl or 0.1N NaOH Prepare fresh Collagenase/dispase solution 1 U/ml collagenase type V (Sigma cat. no. C9263) 0.5 U/ml dispase II (protease type IX, Sigma cat. no. P6141) PBS (Sigma cat. no. D8662), sterile Depending on enzyme activity (U/weight), weigh appropriate amounts to prepare a 50-ml stock in PBS. Filter through a 0.22-µm syringe filter and store in 10-ml aliquots up to 6 months at −20◦ C.

Collagen solution, 1 mg/ml Prepare a 1 mg/ml collagen type I (Sigma cat. no. C9791) solution with a final 20% glacial acetic acid (Merck cat. no. 1.00063) concentration in distilled water. Transfer 250 mg of lyophilized collagen type 1 to a sterile glass bottle. Gradually add 50 ml of glacial acetic acid. Due to variable purity in different collagen preparations, the time necessary for complete dissolution of collagen may vary. Overnight incubation at room temperature is recommended. After complete collagen dissolution in acetic acid, gradually add 200 ml of distilled water. Mix gently without shaking. Store up to 6 months at 4◦ C. To obtain an efficient solution, it is very important to wait for the collagen to be completely dissolved in acetic acid before adding distilled water.

D2 medium High-glucose DMEM (Sigma cat. no. S8636) supplemented with: 2% (v/v) horse serum (heat-inactivated HS) 2 mM glutamine (Sigma cat. no. 67513) Penicillin-streptomycin (10,000 U/ml and 10 µg/ml, respectively; Sigma cat. no. P0781) 1 mM sodium pyruvate (Sigma cat. no. S8636) Store up to 4 weeks at 4◦ C D10 medium Hihg-glucose DMEM (Sigma cat. no. D5671) supplemented with: 10% (v/v) fetal bovine serum (heat-inactivated FBS) 2 mM glutamine (Sigma cat. no. 67513) Penicillin-streptomycin (10,000 U/ml and 10 µg/ml, respectively; Sigma cat. no. P0781) 1 mM sodium pyruvate (Sigma cat. no. S8636) Store up to 4 weeks at 4◦ C

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D20 medium High-glucose DMEM (Sigma cat. no. D5671) supplemented with: 20% (v/v) fetal bovine serum (heat-inactivated FBS) 2 mM glutamine (Sigma cat. no. 67513) Penicillin-streptomycin (10,000 U/ml and 10 µg/ml, respectively; Sigma cat. no. P0781) 1 mM sodium pyruvate (Sigma cat. no. S8636) Store up to 4 weeks at 4◦ C Freezing solution Prepare a mixture of 90% (v/v) FBS and 10% (v/v) DMSO (Sigma cat. no. D2650). Prepare fresh and store up to 24 hr at 4◦ C.

Human bFGF stock solution, 50 µg/ml Reconstitute 50 µg human basic FGF (Peprotech cat. no. 100-18B) in 1 ml of 10 mM Tris·Cl, pH 7.6. Store in 25-µl aliquots up to 6 months at −20◦ C.

M5 medium Megacell DMEM (Sigma cat. no. M3942) 5% (v/v) fetal bovine serum (heat-inactivated FBS) 2 mM glutamine (Sigma cat. no. 67513) 0.1 mM β-mercaptoethanol (GIBCO cat. no. 31350-010) 1% (v/v) non-essential amino acids (Sigma cat. no. M7145) 5 ng/ml human bFGF (Peprotech cat. no. 100-18B) Penicillin-streptomycin (10,000 U/ml and 10 µg/ml, respectively; Sigma cat. no. P0781) Store up to 2 weeks at 4◦ C Matrigel stock Thaw a 10-ml bottle of growth factors–reduced Matrigel (BD Biosciences cat. no. 356230) overnight on ice. Prepare aliquots with sterile microcentrifuge tubes chilled on ice and pipet tips kept at 4◦ C. Store undiluted Matrigel in 100-µl aliquots up to 12 months at −20◦ C. Concentrated Matrigel solution tends to polymerize very quickly at room temperature. Preparation of aliquots must be carried out carefully on ice. Matrigel matrix (BD Biosciences) is a soluble basement membrane extract of the Engelbreth-Holm-Swam (EHS) tumor that gels at room temperature to form a genuine reconstituted basement membrane. The major components of matrigel matrix are laminin, collagen IV, entactin, and heparan sulfate proteoglycan. Growth factors, collagenases, plasminogen activators, and other undefined components have also been reported in the matrigel matrix. The concentration of each component is reported in the product specification sheet.

Mitomycin C Reconstitute a 2-mg ampule of mitomycin powder (Sigma cat. no. M0503) in 1 ml of sterile PBS. Store stock solution (1 mg/ml), protected from light, up to 2 weeks at 4◦ C. Isolation and Characterization of Mesangioblasts

CAUTION: Mitomycin C is a very toxic substance. Handle carefully according to the manufacturer’s product data sheet instructions.

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Oil-Red O solution Weigh 350 mg of Oil-Red O powder (Sigma cat. no. O-0625) and add to 100 ml of 2-propanol (Merck cat. no. 1.09634) in a glass bottle. Let stand overnight at room temperature, protected from light. Do not mix. Filter on 3 MM chromatography paper into a new glass bottle. Add 75 ml of distilled water. Let stand overnight at 4◦ C, protected from light. Do not mix. Filter two times through 3 MM chromatography paper into a new glass bottle. Store up to 6 months at room temperature, protected from light. Oil-Red-O is a very strong staining agent and should be handled carefully according to the manufacturer’s product data sheet instructions.

COMMENTARY Background Information When searching for the origin of the bone marrow cells that contribute to muscle regeneration (Ferrari et al., 1998), a progenitor cell derived from the embryonic aorta has been identified by clonal analysis (De Angelis et al., 1999). When expanded on a feeder layer of embryonic fibroblasts, the clonal progeny of a single cell from the mouse dorsal aorta acquires unlimited lifespan, expresses angioblastic markers (CD34, Sca1, and Flk1), and maintains multipotency in culture or when transplanted into a chick embryo. It is proposed that these newly identified, vessel-associated stem cells, the mesoangioblasts, participate in post-embryonic development of the mesoderm and the authors speculate that post-natal mesodermal stem cells may be rooted in a vascular developmental origin (Minasi et al., 2002). In as much as mesoangioblasts can be expanded in culture, are able to circulate and are easily transduced with lentiviral vectors, they appeared as a potential novel strategy for the cell therapy of genetic diseases. To this aim, it was necessary to isolate mesoangioblast-like cells also from post-natal mouse, dog, and human tissues. This was recently accomplished in the authors’ laboratory. When injected into the blood circulation, mesoangioblasts accumulate in the first capillary filter they encounter and are able to migrate outside the vessel, but only in the presence of inflammation, as in the case of dystrophic muscle. Therefore, it has been reasoned that if these cells were injected into an artery, they would accumulate into the capillary filter and from there into the interstitial tissue of downstream muscles. Intra-arterial delivery of wild-type mesoangioblasts in the αα-sarcoglycan null mouse, a model for limb girdle muscular dystrophy, corrects morphologically and functionally the dystrophic phe-

notype of all the muscles downstream of the injected vessel. Furthermore, mesoangioblasts, isolated from α-sarcoglycan null mice and transduced with a lentiviral vector expressing αα-sarcoglycan, reconstituted skeletal muscle similarly to wild-type cells (Sampaolesi et al., 2003). These data represented the first successful attempt to treat a murine model of muscular dystrophy with a novel class of mesoderm stem cells. To move towards clinical experimentation, canine mesoangioblasts have been isolated. The only animal model specifically reproducing the full spectrum of human pathology is the golden retriever dog model. Affected animals possess a single mutation in intron 6 of the dystrophin gene, resulting in complete absence of the dystrophin protein, and early and severe muscle degeneration with nearly complete loss of motility and walking ability. Intraarterial delivery of wild-type canine mesoangioblasts (vessel-associated stem cells) results in an extensive recovery of dystrophin expression, normal muscle morphology, and function (confirmed by measurement of contraction force on single fibers). The outcome was a remarkable clinical amelioration and preservation of active motility (Sampaolesi et al., 2006). Overall, the data thus far accumulated qualify the mesoangioblasts as candidates for future stem cell therapy for Duchenne patients. Finally, the corresponding human cells were isolated from muscle biopsies. The availability of a large number of human-specific antibodies allowed a complete characterization of these cells, not possible in the corresponding canine cells because of the few available reagents. Data recently published indicated that, in contrast to mouse embryonic mesoangioblasts, human cells, despite a similar morphology and proliferation ability, express pericyte and not endothelial markers, and have therefore been defined as pericyte-derived cells. Human pericyte-derived cells have been shown

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to colonize muscle of dystrophic, immune deficient mice (mdx/SCID) and to give rise to muscle fibers expressing human dystrophin (Dellavalle et al., 2007). A complete understanding of the origin, phenotypic heterogeneity and lineage relationships of this group of cells, also in relationship to other recently described mesoderm stem/progenitor cell, will require further experimental work.

Critical Parameters

Isolation and Characterization of Mesangioblasts

All tissue culture procedures must be performed under strict aseptic conditions. Particular care should be taken to avoid mycoplasma contamination of cell cultures. A sensitive test for mycoplasma detection should be regularly performed (i.e., weekly). Mycoplasma-contaminated cultures should be immediately discarded, or specifically treated in different incubators, if possible, in a different tissue culture room, if they are for some reason irreplaceable. Dissection of mouse embryo aorta must be performed in the shortest possible time due to reduced cell viability with increased dissection time and prolonged tissue manipulation. Aorta fragments should never dry out during dissection and culture. Use of ad hoc humidified chambers in incubators is essential. Do not proceed in protocol steps if primary mesoangioblast outgrowth is not clearly distinguishable (see Basic Protocol 1, step 10). Explants may be cultured for 1 or 2 additional days, but after this period, they should be discarded if a clear cell outgrowth is still undetectable. Collagenase/dispase digestion may be aggressive for the cells, particularly embryonic mesoangioblasts. It is advisable to set the conditions to cell survival rate (see note to Basic Protocol 1, step 20). Each new batch of collagenase and dispase should be tested, since activity of these enzymes may vary between different batches. Some strains of cells currently used as a feeder layer may differ in their abilities to support mesoangioblast clone outgrowth. STO usually support more efficient cloning than MEFs. Different strains of STO and MEF may exhibit different sensitivity to mitomycin C (MMC) inactivation. A titration should be performed to determine the effective MMC dose, both in presence and absence of FBS in culture medium used for MMC inactivation (see Support Protocol 1). An efficient inactivation of the feeder layer represents a crucial point in the cloning proce-

dure. Effective inactivation should be carefully checked setting appropriate controls. Since the feeder layer represents an essential part of the cloning protocol, particular care should be taken in culturing the selected strain of cells. The cells can be used only if they proliferate well and look healthy. Over-confluence cultures must always be avoided. MEFs cannot be used beyond six to eight passages. Special attention must be paid to cell density: initiating cultures at ∼20% confluence (∼1:4 split from an 80% confluent flask) and growing to ∼80% confluence seems to guarantee best efficiency in cell proliferation and preservation of differentiation capability. Excessive dilution may result in growth arrest. Over-confluence, leading to acidification of culture medium, may cause uncontrolled spontaneous differentiation. Operationally, it is recommended to (1) freeze newly derived mesoangioblasts at very early passage; (2) periodically check differentiation ability (see below); and (3) periodically perform karyotype analysis. Physiological O2 tension is essential for mesoangioblast cultures. The traditional cell culture gas mix, consisting of 5% CO2 in air, contains oxygen ranging from 18% to 21%. These conditions represent a hostile environment, since the concentration of oxygen in most mammalian tissue is equivalent to 3% to 5%. Therefore, excess oxygen can result in the generation of reactive oxygen species leading to DNA damage, chromosomal instability, and stasis. Growth curves for human pericytederived cells are shown in Figure 2B.1.2, comparing 5% O2 and air O2 tension. The use of low oxygen seems to be particularly important in cloning experiments. Cloning is a very stressful event for most normal diploid cells. Although a high fraction of low-passage cells divide when sub-cultivated under normal conditions, cloning efficiencies (growth of attached cells) are typically only 1% to 10%, in spite of high-plating efficiency (simple survival of attached cells). This is due to many diverse variables, including oxygen toxicity (Wright and Shay, 2002). For murine mesoangioblasts, usually cloned on a murine feeder layer, the use of a low-oxygen environment represents an added strategy to obtain higher cloning efficiency. For human pericyte-derived cells, the use of this system allows the isolation of proliferating clones without the use of any murine feeder layer. The effect of physiological oxygen tension on cell culture can be studied by any laboratory. An inexpensive

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Figure 2B.1.2 Proliferation curve of human pericyte-derived cells under low (5%, diamonds) and high (20%, squares) O2 concentration.

low-oxygen incubator can be produced from commercially available, simple gas-tight containers that can be flushed with prepared gas mixtures to produce low-oxygen environments for test cell cultures (Wright and Shay, 2006). A common problem observed when culturing adult human cells are chromosome rearrangements and proliferation arrest. Most mammalian normal cells do not divide indefinitely, owing to replicative senescence. In human cells, replicative senescence is caused by telomere shortening, even in cells with detectable telomerase activity. When tested for telomerase activity, human mesoangioblasts at early passage (VIII) show a significant TRAP activity (5% to 10% of that found in reference cancer cells). However, at later passages (XIX), telomerase activity is no longer detectable. Consistently, telomere length progressively shortens, reaching a size typical of pre-senescent cells. Telomere shortening is thought to induce senescence through the activation of DNA damage signals (p53-mediated pathway). Senescence, however, may be caused in culture also by telomerase-independent oxidative stress. This mechanism (p16 mediated) is defined as stasis (stimuli- and stress-induced senesce like growth arrest; Wright and Shay, 2002) and represents a main issue in in vitro culturing of cells, including murine cells.

dishes, mesoangioblasts acquire their definitive morphology of adherent cells and frequently undergo a critical phase. Since high density of cells during this phase is crucial to their survival; growth arrest may be due to excessive cell dilution. Collecting and replating cells at higher density may help to recover healthy clones. Trypsinization during these early, critical phases should be particularly mild (3 min at 37◦ C), avoiding hard trituration of cell pellet during resuspension. Trypsin has to be completely removed and the cell pellet suspended in freshly prepared, prewarmed medium. 2. Isolated clones do not detach. Difficulty in trypsinization may be due to: early mycoplasma contamination of cells. An appropriate detection test has to be performed immediately. Spontaneous differentiation of the culture: in this case a change in morphology should progressively appear, mainly to adipocytes or smooth muscle cells. These clones must be discarded. 3. Very low cloning efficiency and/or high rate in spontaneous differentiation. Mesoangioblast isolation may be difficult in some mouse strains (i.e., Balb/C). In this case, it is advisable to expand cells as a polyclonal mix first. Afterwards, appropriate cell cloning or sorting can be performed.

Troubleshooting

Dog and human postnatal skeletal muscle 1. Poor primary mesoangioblast outgrowth from muscle explant: Particularly in human samples, a high variability in proliferation of primary cells can be observed. The isolation of pericyte-derived cells may be difficult if the round shaped, refractile cell population

Mouse embryo aorta and postnatal skeletal muscle 1. Isolated clones drastically reduce or arrest proliferation after first/second passage. After the first passage onto uncoated plastic

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described is not easily distinguishable, or if just a few floating cells are present. In this case it is advisable to expand (for a single passage) the whole polyclonal cell mix, proceeding as follows: (a) Remove culture medium and set it aside in a 15-ml centrifuge tube. (b) Carefully detach and discard muscle fragments using a 1000-µl pipettor. (c) Rinse the growing surface with 2 to 5 ml of CMF-PBS (depending on dish size). Add 2 to 5 ml of trypsin and incubate 5 min at 37◦ C, monitoring the complete detachment of all cells under a microscope. (d) Use the saved culture medium to collect cells. Centrifuge 10 min at 232 × g. Suspend accurately the pellet in freshly prepared, prewarmed medium and plate cells in petri dish (same size of dish used for primary culture assembling). (e) Incubate 1 to 3 days in a 37◦ C, 5% CO2 incubator. At this point, the floating population of pericyte-derived cells should be easily distinguishable. Transfer medium and floating cells to a new dish or 25-cm2 tissue culture flask. Discard the primary mixed population of adherent cells. 2. Early or intermediate passage cells (up to passage 10) stop growing. If cells do not need to be split within 3 to 4 days, they probably have been diluted too much. In this case, even if not yet reaching optimal density, cells have to be detached, and plated to a higher density. Monitor cell proliferation in the following 48 hr. If cells do not start to proliferate again regularly, they should be discarded.

Isolation and Characterization of Mesangioblasts

Myogenic differentiation 1. Cells do not differentiate in the coculture assay: Co-culture with mouse and rat myoblasts is the standard procedure to test myogenic differentiation of mesoangioblasts. Thus, it is crucial that inducer strains of rodent myoblasts such as C2C12 and L6, are properly maintained and checked for their myogenic differentiation. In particular, cell density should never be >70% or <20%. At high density, myogenic cells differentiate also in proliferation-inducing media; in contrast, at low density clones develop, cells reach a local high density (within the clone) and also begin to differentiate. In both cases, cells less prone to differentiation are then subcultured and with time the whole population may become differentiation deficient. It is crucial to frequently check myogenic differentiation of inducer myoblasts.

Anticipated Results Cloning of mouse embryo aorta mesoangioblasts has an efficiency of ∼5% to 10%. The cloning efficiency decreases if primary source is mouse, dog, or human adult tissue. Cloning efficiency may vary depending on culture conditions (e.g., O2 level), mouse strains (Balb C, for example, give very few mesoangioblast clones), and type of feeder cell (STO usually result in higher cloning efficiency than MEF). High variability in cell proliferation rate is observed in different dog and human primary mesoangioblast cultures derived from skeletal muscle, independent of age. The presence of fat infiltration in dog and human skeletal muscle biopsies is very frequently associated to a slower mesoangioblast outgrowth in culture. Once established, mesoangioblasts clones and/or polyclonal cell mix can be expanded efficiently, but not indefinitely. Senescence can be expected at or near passage 30 for mouse-derived cells and passage 20 for dog- and human-derived cells. Different factors, however, may significantly affect mesoangioblast replicative lifespan and integrity: culture conditions (O2 level); appropriate media supplements (human pericytederived cells need bFGF and reducing agents such as 2β-mercaptoethanol); care in propagation, splitting and freezing—an average density of 20% to 80% should be maintained. At every freeze/thaw cycle, 10% to 20% cell death should be expected.

Time Considerations Supplementing and filtering of media requires ∼1.5 hr, including FBS inactivation (45 min). Preparation of the feeder layer Mitomycin cell inactivation requires 3 hr. Counting, diluting, and plating inactivated cells onto 48-well plates to be used in a cloning experiment (twelve plates) requires ∼1.5 hr. Collagen coating The preparation of collagen solution is time consuming and may vary depending on collagen purity. An overnight incubation is advisable, even if 3 to 4 hr may be sufficient for collagen to dissolve. Time necessary for collagen coating of plastic dishes depends mainly on the fact that collagen-treated dishes must be completely dried in a dediccated oven overnight at 30◦ C. Matrigel coating Time necessary for Matrigel coating of dishes depends on incubation time for Matrigel

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polymerization (30 min), and on the total number of plates to be coated, since, as mentioned above, prior to coating it is essential to keep all reagents on ice and thus only a few dishes should be coated at the same time. Dissection of mouse embryo aorta The time needed to dissect the embryonic aorta depends upon the experience of the investigator and the number of embryos to be processed (from 0.5 to 2 hr). It is advisable to start with a small number of embryos and not to keep dissected aortas under the hood for more than 2 hr. Primary culture: dissection of adult skeletal muscle Isolation of individual muscles, dissection, and mincing the tissue into fragments requires ∼30 to 45 min (for 100 to 200 mg skeletal muscle sample). Collagenase/dispase digestion requires ∼15 to 20 min for sample preparation and 15 min of incubation. Cloning experiment For mouse mesoangioblasts, accurate counting, preparation of four different dilutions, and the final plating onto inactivated feeder-coated multiwell plates of the cells requires ∼1.5 to 2 hr (twelve plates). For human mesoangioblasts (pericyte-derived cells), cloning is carried out onto collagen-coated multiwell plates. The time necessary for ten plates is ∼1 hr. Freezing/Thawing Freezing/thawing of mouse, dog, and human mesoangioblasts requires ∼30 min.

Literature Cited Cepko, C. 1996. Preparation of a specific retrovirus producer cell line. Curr. Protoc. Molec. Biol. 136:9.10.1-9.10.13. Chaplin, A.J. and Grace, S.R. 1975. Calcium oxalate and the von Kossa method with reference to the influence of citric acid. Histochem. J. 7:451458. Davidson, J.G. 1998. Immunofluorescent staining. Curr. Protoc. Cell Biol. 0:4.3.1-4.3.6. De Angelis, L., Berghella, L., Coletta, M., Lattanzi, L., Zanchi, M., Cusella-De Angelis, M.G., Ponzetto, C., and Cossu, G. 1999. Skeletal myogenic progenitors originating from embryonic dorsal aorta coexpress endothelial and myogenic markers and contribute to postnatal muscle growth and regeneration. J. Cell Biol. 147:869878.

Dellavalle, A., Sampaolesi, M., Tonlorenzi, R., Tagliafico, E., Sacchetti, B., Perani, L., Innocenzi, B., Galvez, B.G., Messina, G., Morosetti, R., Li, S., Belicchi, M., Peretti, G., Chamberlain, J.S., Wright, W.E., Torrente, Y., Ferrari, S., Bianco, P., and Cossu, G. 2007. Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat. Cell Biol. 9:255-267. Ferrari, G., Cusella-De Angelis, G., Coletta, M., Paolucci, E., Stornaiuolo, A., Cossu, G., and Mavilio, F. 1998. Muscle regeneration by bone marrow-derived myogenic progenitors. Science 279:1528-1530. Hogan, B., Beddington, R., Costantini, F., and Lacy, E. 1994. Section C. Recovery, culture, and transfer of embryos and cells. In Manipulating the Mouse Embryo (2nd Edition) pp. 167-168. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Lin, M.S., Alfi, O.S., and Donnell, G.N. 1976. Differential fluorescence of sister chromatids with 4 -6-diamidino-2-phenylindole. Can J. Genet. Cytol. 18:545-547. Minasi, M.G., Riminucci, M., De Angelis, L., Borello, U., Berarducci, B., Innocenzi, A., Caprioli, A., Sirabella, D., Baiocchi, M., De Maria, R., Jaffredo, T., Broccoli, V., Bianco, P., and Cossu, G. 2002. The meso-angioblast: A multipotent, self-renewing cell that originates from the dorsal aorta and differentiates into most mesodermal tissues. Development 129:27732783. Ross, J.J., Hong, Z., Willenbring, B., Zeng, L., Isenberg, B., Lee, E.H., Reyes, M., Keirstead, S.A., Weik, E.H., Tranquillo, R.T., and Verfaillie, C.M. 2007. Cytokine-induced differentiation of multipotent adult progenitor cells into functional smooth muscle cells. J. Clin. Invest. 116:31393149. Sampaolesi, M., Torrente, Y., Innocenzi, A., Tonlorenzi, R., D’Antona, G., Pellegrino, M.A., Barresi, R., Bresolin, N., Cusella-De Angelis, M.G., Campbell, K.P., Bottinelli, R., and Cossu, G. 2003. Cell therapy of alpha sarcoglycan null dystrophic mice through intra-arterial delivery of mesoangioblasts. Science 301:487-492. Sampaolesi, M., Blot, S., D’Antona, G., Granger, N., Tonlorenzi, R., Innocenzi, A., Mognol, P., Thibaud, J.L., Galvez, B.G., Barth´el´emy, I., Perani, L., Mantero, S., Guttinger, M., Pansarasa, O., Rinaldi, C., Cusella De Angelis, M.G., Torrente, Y., Bordignon, C., Bottinelli, R., and Cossu, G. 2006. Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature 444:574-579. Wright, W.E. and Shay, J.W. 2002. Historical claims and current interpretations of replicative aging. Nat. Biotechnol. 20:682-688. Wright, W.E. and Shay, J.W. 2006. Inexpensive low-oxygen incubators. Nat. Protoc. 1:20882090. Somatic Stem Cells

2B.1.29 Current Protocols in Stem Cell Biology

Supplement 3

Purification and Culture of Human Blood Vessel–Associated Progenitor Cells

UNIT 2B.2

Mihaela Crisan,1, 4 Johnny Huard,1, 2, 4 Bo Zheng,3, 4 Bin Sun,1, 4 Solomon Yap,1, 4, 5 Alison Logar,3, 4 Jean-Paul Giacobino,1, 3, 4 Louis Casteilla,1, 6 and Bruno P´eault1, 2, 4 1

Children’s Hospital of Pittsburgh of UPMC, Pittsburgh, Pennsylvania McGowan Institute for Regenerative Medicine, Pittsburgh, Pennsylvania 3 Department of Orthopedic Surgery and Molecular Genetics and Biochemistry, Children’s Hospital of Pittsburgh of UPMC, Pittsburgh, Pennsylvania 4 Stem Cell Research Center, Children’s Hospital of Pittsburgh of UPMC, Pittsburgh, Pennsylvania 5 University of Pittsburgh, Pittsburgh, Pennsylvania 6 University of Toulouse, Toulouse, France 2

ABSTRACT Multilineage progenitor cells, diversely designated as MSC, MAPC, or MDSC, have been previously extracted from long-term cultures of fetal and adult organs (e.g., bone marrow, brain, lung, pancreas, muscle, adipose tissue, and several others). The identity and location, within native tissues, of these elusive stem cells are described here. Subsets of endothelial cells and pericytes, which participate in the architecture of human blood vessels, exhibit, following purification to homogeneity, developmental multipotency. The selection from human tissues, by flow cytometry using combinations of positive and negative cell surface markers, of endothelial and perivascular cells is described here. In addition, a rare subset of myoendothelial cells that express markers of both endothelial and myogenic cell lineages and exhibit dramatic myogenic and cardiomyogenic potential has been identified and purified from skeletal muscle. The culture conditions amenable to the long-term proliferation of these blood vessel–associated stem cells in vitro are C 2008 by John Wiley also described. Curr. Protoc. Stem Cell Biol. 4:2B.2.1-2B.2.13.
& Sons, Inc. Keywords: endothelial cell r myogenic cell r blood vessel r pericyte r flow cytometry

INTRODUCTION The prospective isolation by flow cytometry of different cell subsets that build up or are developmentally affiliated with the walls of human blood vessels is described here. Subsets of perivascular cells, i.e., cells that immediately surround endothelial cells in blood vessels, including pericytes, which enwrap endothelial cells in capillaries and microvessels, and a novel population of skeletal muscle myoendothelial cells, that co-express markers of both myogenic and endothelial cells and have the potential to differentiate into skeletal and cardiac myofibers, bone, fat, and cartilage cells, are presented. Protocols used for the procurement, storage, and handling of human fetal and adult tissues (see Support Protocol 1), the enzymatic dissociation and antibody staining of human tissues (see Basic Protocol 1), the purification by fluorescence activated cell sorting (FACS) of discrete vascular cell subsets (see Basic Protocol 2), and the assessment of sorted cell purity by reverse transcription-polymerase chain reaction (RT-PCR; see Support Protocol 2) are all described in this unit. In addition, the conditions used for the long-term culture of these cells, in which their developmental potential is sustained, is described (see Basic Protocol 3). Somatic Stem Cells Current Protocols in Stem Cell Biology 2B.2.1-2B.2.13 Published online March 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02b02s4 C 2008 John Wiley & Sons, Inc. Copyright


2B.2.1 Supplement 4

BASIC PROTOCOL 1

PREPARATION OF MYOENDOTHELIAL CELLS AND PERICYTES FROM FETAL AND ADULT TISSUES Against the common belief that the same simple enzyme treatment can be used to dissociate all tissues, dissociation protocols must be refined to efficiently separate endothelial cells from perivascular cells. Numerous variations have been tested regarding the type of enzyme used, and the duration and temperature of incubation. The method described below has been found to work efficiently. After the single-cell suspension has been prepared, pericytes are purified as CD146+ CD34− CD45− CD56− cells.

Materials Fresh embryonic, fetal, or adult tissue (see Support Protocol 1) 100 µg/ml type-I collagenase (Invitrogen) 100 µg/ml type-IV collagenase (Invitrogen) 1.2 µg/ml dispase (Invitrogen) 1× Hanks’ balanced salt solution (HBSS; Invitrogen) 1× Dulbecco’s modified Eagle medium (high-glucose DMEM, Invitrogen) Fetal bovine serum (FBS, Invitrogen) 1% penicillin/streptomycin solution (PS, Invitrogen) Mouse serum (Sigma) 1× Dulbecco’s phosphate buffered saline without calcium and magnesium (CMF-DPBS; Invitrogen) Antibodies: CD45-FITC (BD Biosciences) or CD45-APC-Cy7 (1:100; Santa Cruz Biotechnologies) CD144-PE (Beckman Coulter) CD146-FITC (1:100; Serotec) UEA-1-PE (Biomeda Corporation) CD34-APC (BD Biosciences) or CD34-PE (1:100; DAKO or BD Biosciences) CD56-PE-Cy7 (1:100; BD Biosciences or Serotec) or CD56-APC (BD Biosciences) Isotype control antibodies: APC-Cy7-, APC-, PE-Cy7- and PE-mouse IgG1 (BD Biosciences); PE- (1:100; Chemicon), APC-Cy7- (1:100; BD Biosciences), PE-Cy7- and FITC-conjugated mouse IgG (1:100; US Biological) 7-amino-actinomycin D (7-AAD, ViaProbe; BD Biosciences) CompBeads set (BD Biosciences) Collagenases type IA-S, II-S, and IV-S (Sigma) Trypsin-EDTA (Invitrogen) Trypan blue (Sigma)

Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

37◦ C rotator Sterile surgery scissors, forceps (VWR) and scalpels (Bard-Parker) 50-ml tubes (Falcon) 40-, 70-, and 100-µm cell strainers (BD Biosciences) Flow cytometer CellQuest software (BD Biosciences) 5-ml FACS tubes (BD Biosciences) 60 × 15–mm Petri dishes (BD Biosciences) 37◦ C shaker Polylysine-coated glass slides (ESCO) Hemacytometer Vortex agitator Additional reagents and equipment for cell counting (UNIT 1C.3)

2B.2.2 Supplement 4

Current Protocols in Stem Cell Biology

To purify myoendothelial cells 1a. Carefully remove connective tissue and fat from the adult muscle biopsy, using forceps and scissors. 2a. Cut the muscle biopsy into small pieces, then finely mince with scissors and digest for 60 min at 37◦ C on a rotator (20 rpm) with the following enzyme mixture: 100 µg/ml type-I collagenase, 100 µg/ml type-IV collagenase, and 1.2 µg/ml dispase in HBSS in a 50-ml tube. Usually, 1 g of tissue is digested in 10 ml enzyme solution. Adult tissue samples are usually in the 5- to 10-g range.

3a. Pellet the digested tissue by centrifuging 5 min at 500 × g, 4◦ C. Discard the supernatant. 4a. Resuspend pellet in 10 ml DMEM/10% FBS/1% penicillin/streptomycin, then pass through a 40-µm strainer to obtain a single-cell suspension at room temperature. 5a. For flow cytometry analysis of endothelial, myogenic, and myoendothelial cells, incubate ∼1.0 × 105 dissociated cells 10 min in 50 µl mouse serum diluted 1:10 in CMF-DPBS on ice. 6a. Incubate cells 30 min on ice with 0.1 ml of appropriately diluted (in CMF-DPBS, 2% FBS) mouse monoclonal antibody to human CD45-FITC, CD34-PE, CD56-APC, CD144-PE, CD146-FITC, or with UEA-1-PE. When multiple antibody staining is used (CD45-FITC, CD34-PE, CD56-APC), each antibody is used individually and cells are washed by centrifuging 5 min at 500 × g, 4◦ C, prior to adding 0.1 ml of the next antibody. Negative control samples receive the same amounts of FITC-, APC-, and PE-conjugated isotype-matched pre-immune antibodies.

7a. Analyze a minimum of 1 × 105 live cells by flow cytometry, using the CellQuest software. Typical results are presented in Table 2B.2.1.

8a. For cell sorting, resuspend 5–6 × 106 cells in 500 µl to 1 ml of DMEM/2% FBS in a 5-ml FACS tube. Incubate simultaneously with 0.1 ml of each antibody: APC-Cy7conjugated mouse anti-human CD45, APC-conjugated mouse anti-human CD34, PE-Cy7-conjugated mouse anti-human CD56, PE-conjugated mouse anti-human CD144, or with matching control antibodies. Add 20 µl of 7-AAD for each 1 × 106 cells to each tube for dead cell exclusion. Evaluate background staining with isotype-matched control antibodies and use a CompBeads set to optimize fluorescence compensation settings for multi-color analyses and sorts. Usually, ∼7 × 105 cells are recovered per gram of adult muscle and fetal muscle can yield ten times more cells than adult muscle. For FACS analysis, cultured cells are first detached from flasks with 0.1% trypsin/EDTA, washed in cold PBS/2% FBS, and then incubated with antibodies as described above. Typical results are presented in Table 2B.2.2. Table 2B.2.1 Marker Expression in Human Skeletal Muscle as Percent Positive Cells (n = 11)

Marker Average ± SEM

CD45

CD34

CD144

CD56

UEA-1

CD146

1.02 ± 0.36

14.31 ± 3.74

6.2 ± 2.57

5.79 ± 1.64

51.24 ± 3.79

6.78 ± 2.14

Somatic Stem Cells

2B.2.3 Current Protocols in Stem Cell Biology

Supplement 4

Table 2B.2.2 Cell Subsets Sorted from Human Skeletal Muscle (n = 13)

Cell type

Myogenic

Endothelial

Myoendothelial

Marker

CD56+ CD34− CD144− CD45−

CD56− CD34+ CD144+ CD45−

CD56+ CD34+ CD144+ CD45−

3.0 ± 0.7

5.0 ± 1.2

0.4 ± 0.1

Number ∼10

4.8 ± 1.3

7.7 ± 2.6

1.5 ± 0.8

% purity

93.8 ± 1.0

92.2 ± 1.1

93.5 ± 1.8

% positive 4

To purify pericytes 1b. Using a scalpel and forceps, cut fetal or adult tissues into small pieces (∼0.5cm pieces) in a 60 × 15–mm Petri dish with enough DMEM/20% FBS/1% PS/0.5 mg/ml of each type of collagenase (IA-S, II-S, and IV-S) to completely cover the pieces. 2b. Transfer the medium and tissue pieces into a 50-ml tube and incubate 75 min on a 37◦ C shaker at 120 rpm. Dissociate fetal brain in the collagenase solution for only 45 min. Dissociate bone marrow from long bones by removing epiphyses and cutting the diaphysis longitudinally to expose the marrow. Immerse the two halves in the collagenase solution in a 50-ml tube and incubate for 45 min at 37◦ C, under agitation. Complete the marrow cell dissociation by pipetting.

3b. Apply the undissociated clumps to a polylysine-coated glass slide, cover with a second ground glass slide, and grind the two slides together to break up the clumps. Then rinse the slides with medium (minus enzymes) to recover the suspension. 4b. Filter the resulting suspension through a 100-µm cell strainer and centrifuge 5 min at 300 × g, 4◦ C. 5b. Resuspend the pellet in 1 ml DMEM/20% FBS/1% PS, avoiding bubbles, and count cells by taking 10 µl of cell suspension and mixing with 10 µl of Trypan blue to label dead cells. Count cells on a hemacytometer (UNIT 1C.3). 6b. Resuspend ∼30 × 106 cells in 1 ml of complete medium, and add 10 µl of each appropriately diluted antibody (CD146, CD34, CD56, and CD45). 7b. Vortex gently, then incubate 15 min at 4◦ C in the dark. Incubate 3 × 104 cells in the same conditions with isotype control antibodies. 8b. Wash cells by centrifuging 5 min at 300 × g, 4◦ C. 9b. Resuspend each pellet in 3 ml of DMEM/20% FBS/1% PS and transfer into 5-ml FACS tubes. Resuspend pellet incubated with the isotype control antibody in 500 µl of the same medium in another FACS tube. 10b. Incubate cells 30 min at room temperature with 30 µl 7-AAD (1:100) for dead cell exclusion. Filter the cell suspension on a 70-µm cell strainer prior to sorting. SUPPORT PROTOCOL 1 Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

PROCUREMENT AND STORAGE OF HUMAN FETAL AND ADULT TISSUES Human fetal tissues are obtained from spontaneous, elective, or medical pregnancy interruptions, with the informed consent of the donor and in compliance with the rules established by the Institutional Ethics Committee. Adult tissues are obtained from surgery, upon which they should have normally been discarded, or autopsy. Adult tissues are also collected following rules in effect at the institution/state/country.

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Current Protocols in Stem Cell Biology

NOTE: All studies with human subjects must be approved by the Institutional Review Board (IRB), which must adhere to the Office for the Protection from Research Risk (OPRR) guidelines or other applicable governmental regulations for using human subjects. All material must be obtained with informed consent of the donor.

Materials Fetal or adult tissue samples Hanks’ balanced salt solution (HBSS; Invitrogen) 1× Dulbecco’s phosphate buffered saline without calcium and magnesium (CMF-DPBS; Invitrogen) Fetal bovine serum (FBS, Invitrogen) Penicillin/streptomycin solution (PS, Invitrogen) 1× Dulbecco’s modified Eagle medium (high-glucose DMEM, Invitrogen) Horse serum (Invitrogen) 60 × 15–mm Petri dishes (BD Biosciences) Vertical laminar flow hood Sterile surgery scissors, forceps (VWR), and scalpels (Bard-Parker) Dissecting microscope 1. Place fetal (e.g., muscle, pancreas, bone marrow, brain, lung, and placenta) or adult human tissue samples (muscle, fat, pancreas) in a 60 × 15–mm petri dish containing HBSS or CMF-DPBS, both containing 5% FBS, 1% PS, for transfer to the laboratory. Fetal tissues can be stored for up to 6 days at 4◦ C in high-glucose DMEM/20% FBS/1% PS. Adult muscle is stored preferably for up to 7 days in DMEM/50% horse serum/1% PS at 4◦ C with good cell viability. Rules differ between countries/institutions regarding the serologic tests performed on the donors. The HBV/HIV serology is usually known and the use of any known contaminated human tissue is strongly discouraged. All human tissues should be considered as potentially infectious and processed accordingly. Latex gloves and laboratory coats should be worn at all times and any drinking/eating should be strictly prohibited in the laboratory. Material should be disposed of in the appropriate waste containers for infectious materials and liquid waste should be treated with bleach for at least 10 min before discarded. Reusable containers and instruments must be autoclaved.

2. When necessary, dissect the tissues under a vertical laminar flow hood using sterile surgery scissors and forceps. This may require the use of a dissecting microscope. Do not use any human tissue that has been previously frozen as this would result in poor cell viability. Only use fresh tissues throughout the procedure, preferably as soon after harvesting as possible.

CELL SORTING TO PURIFY HUMAN ENDOTHELIAL, MYOENDOTHELIAL, AND PERIVASCULAR CELLS

BASIC PROTOCOL 2

This protocol describes the purification of human endothelial, myoendothelial, and perivascular cells by multicolor fluorescence-activated cell sorting, using combinations of positive and negative cell-surface markers. Hematopoietic (CD45+ ) cells are gated out in all instances. Endothelial cells are selected on expression of both CD144 (VE-cadherin) and CD34 (CD45− CD144+ CD34+ ). Myoendothelial cells are sorted on additional expression of CD56 (CD45− CD144+ CD34+ CD56+ ). Perivascular cells are purified on expression of CD146 and absence of CD34 (CD146+ CD34− ): since CD146 is also expressed on endothelial cells, negative selection of CD34+ cells allows one to sort pure perivascular cells. Somatic Stem Cells

2B.2.5 Current Protocols in Stem Cell Biology

Supplement 4

Materials Sterile PBS (CMF-DPBS) SPHERO Rainbow fluorescent particles (BD Biosciences) FACS Accudrop fluorescent beads (BD Biosciences) 70% ethanol Antibody stained cell suspensions (see Basic Protocol 1) BD FACSAria flow cytometer equipped with three lasers (488-, 633-, and 407–nm; BD Biosciences) with the following standard filter configurations: Blue excitation: FITC 525 nm, PE 575 nm, PE-Texas Red 610 nm, PerCP-Cy5.5 695 nm, PECy7 780 nm Red excitation: APC 660 nm, APCCy7 780 nm Violet excitation: Pacific Blue 450 nm, Pacific Orange 530 nm 1. Set the FACSAria flow cytometer to sort in high mode. Install a 70-µm tip in the instrument and complete sorting at 70 psi. This allows the detection of nine fluorescent cell subpopulations as well as forward scatter and side scatter.

2. Set frequency and amplitude to yield a steady stream with minimal satellite droplets. Frequency and amplitude for sorting stream set up is specific for the particular nozzle installed in the instrument. These droplets can decrease the purity of the sort by allowing non-desired events to be collected.

3. Set the flow rate at setting 2. Cells are run at a fairly slow flow rate so as not to increase the abort count by running cells at a higher rate. This slower rate also cuts down on clogs as the dissociated tissue will clump and clog the instrument as it sorts. If clumps do occur, samples are filtered through a 70-µm filter prior to resuming the sort.

4. Use sterile PBS as sheath fluid. 5. Align the FACSAria prior to sorting using SPHERO Rainbow fluorescent particles. These particles will show proper alignment of the lasers indicated by sharp peaks on fluorescent histograms as well as proper calculation of the laser delays by comparison of area versus height signals for each laser. Area scaling is set based on the biological sample, not the bead preparation.

6. Determine the proper drop delay by running FACS Accudrop beads.

Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

These beads are run through the instrument and deflected while the drop delay is altered. The drop delay, which yields the highest percent of deflection and capture in the optical filter window, is the delay used for subsequent sorting of the biological sample. The precision setting for sorting of the sample is typically set to Purity 32 Phase 16. This setting determines when a cell will be sorted based on where the cell is in the droplet to be selected and how close contaminating cells are to the desired cell. Each droplet is separated into 32 increments. A purity value of 32 determines that if a contaminating cell is within 32 increments (16 leading and 16 trailing or half a drop above or below) the desired cell will not be collected. The Phase 16 value determines where a desired cell is found within the droplet and whether or not it is selected. The sorted cell must be centered within the middle 16 increments of the droplet to be selected. Therefore, cells close to the edge of the droplet will not be selected. Cells close to the edge can affect the trajectory of the deflection of the cell. A Purity 32 Phase 16 mode will eliminate many cells from being sorted, reducing the recovery of the desired cells, but will yield high purity of the cells sorted.

2B.2.6 Supplement 4

Current Protocols in Stem Cell Biology

7. Sterilize the instrument prior to sorting by running the Prepare for Aseptic Sort program using the FACSAria software. This program runs 70% ethanol through the sheath and sample lines to eliminate contamination of sorted cells.

8. For each sample analysis, initially create an area versus height dotplot for forward scatter as well as a signal specific for each laser. 9. Create a side scatter dotplot for the blue laser, APC for the red laser, and Pacific Blue for the violet laser. 10. Set the area scaling value such that the cells are along the 45◦ angle between the two signals. One signal should not be brighter at the expense of the other. Area scaling is set for each particular cell type. 7-AAD is added to the stained cells prior to sorting to eliminate dead cells from all subsequent analysis and sorting.

11. Create a standard forward scatter versus side scatter dotplot to eliminate debris and clumps of cells from analysis. 12. Use a height versus width dotplot to eliminate doublets within the forward versus side scatter gate. Doublets have an increased cell width and are removed, as they will decrease cell purity if they are included in the analysis.

13. Gate all fluorescent histograms on the viable 7-AAD-negative cells, which are free of debris, clumps, and doublets, based on the light scatter properties of the cells. Use irrelevant fluorescently matched antibodies as negative controls. Use single color–stained cells as compensation controls. 14. Sort endothelial cells as CD45− CD56− CD146− CD34+ cells. Sort perivascular cells (pericytes) as CD45− CD56− CD146+ CD34− cells. Sort myoendothelial cells as CD45− CD56+ CD146+ CD34+ CD144+ cells. Ideally, the cytometer is placed in a custom-built BSL-2 containment hood to ensure safety of the operator. This hood will contain suspected pathogens as these human samples are sorted, prior to the availability of screening tests, in compliance with safety guidelines for the handling of human samples.

LONG-TERM CULTURE OF HUMAN BLOOD VESSEL–ASSOCIATED PROGENITOR CELLS

BASIC PROTOCOL 3

A unique property of human myoendothelial and perivascular cells (i.e., pericytes) is that these cells can be cultured long term (passages 16 to 18; i.e., ∼20 weeks) with no significant loss of developmental potential.

Materials Sorted myoendothelial cells (see Basic Protocol 2) Proliferation medium (see recipe) Trypsin/EDTA (Invitrogen) 1× Dulbecco’s modified Eagle medium (high-glucose DMEM, Invitrogen) Fetal bovine serum (FBS, Invitrogen) Penicillin/streptomycin solution (PS, Invitrogen) 0.2% gelatin (Calbiochem) Sorted pericytes (see Basic Protocol 2)

Somatic Stem Cells

2B.2.7 Current Protocols in Stem Cell Biology

Supplement 4

Endothelial cell growth medium (EGM2, Cambrex) DPBS Dimethylsulfoxide (DMSO, Sigma) Liquid nitrogen 6-, 12-, 48-, and 96-well collagen-coated plates (Corning) 37◦ C, 5% CO2 cell incubator Refrigerated low-speed centrifuge 25- and 75-cm2 tissue culture flasks (Falcon) 12- and 48-well uncoated plates 15-ml conical tubes (Falcon) To prepare myoendothelial cell culture 1a. Plate FACS-sorted myoendothelial CD56+ CD34+ CD144+ CD45− cells in 96-well collagen-coated plates at a density of 500 cells per well in 100 µl proliferation medium and incubate in a 37◦ C, 5% CO2 incubator. 2a. At 60% confluence, detach cells by adding 100 µl of 0.1% trypsin/EDTA to each well. Collect the detached cells in 100 µl DMEM, 10% FBS, 1% PS with a pipet and transfer into a 15-ml conical tube. 3a. Wash the cells with 5-ml DMEM/10% FBS/1% PS and centrifuge 5 min at 500 × g, 4◦ C. 4a. Resuspend cell pellet in 350 µl proliferation medium. 5a. Replate the cells on 48-well collagen-coated plates containing 350 µl proliferation medium at densities between 1.0 and 2.5 × 103 /cm2 . 6a. Continue to feed cells with proliferation medium every 2 to 3 days and passage the cells at 5 to 7 days when they reach 60% confluency. Expand cells at each passage to 24-well, 6-well collagen-coated plates and then to 25-cm2 and 75-cm2 tissue culture flasks. Culture further for 3 to 4 weeks to expand cells to have enough for myogenic differentiation and cell transplantation experiments. Cells should exhibit no sign of differentiation during this expansion phase. Test the ability of sorted cells to differentiate into myotubes in vitro by plating them at 1.0 × 105 cells per well in 6-well plates for 2 days in 2 ml proliferation medium. Replace the medium with 2 ml fusion medium (DMEM/2% FBS/1% PS). After 10 to 14 days, cells come to confluence and differentiate into multinucleated myotubes. For cell transplantation experiments, resuspend 3–5 × 105 (cell number) cultured cells mixed with 1µl of 1:1000 FluoSpheres (Molecular Probes) in 15 µl HBSS and transplant in a single injection into a SCID mouse gastrocnemius muscle that was injured 1 day earlier by intramuscular injection of 1 µg cardiotoxin (CTX, Molecular Probes) in 20 µl HBSS. Sacrifice animals 10 days post-injection and harvest treated muscles for chimerism analysis, using human-specific antibodies on tissue sections.

To prepare pericyte cell culture 1b. Treat wells from a 48-well culture plate with 0.2% gelatin for 10 min at 4◦ C. 2b. Culture 2 × 104 sorted pericytes per well in 1 ml EGM2 medium in a 37◦ C, 5% CO2 incubator. Change EGM2 medium after 7 days, then every 4 days until 100% confluency is reached (∼2 weeks). Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

Freshly sorted pericytes attach after 3 to 4 days of culture, and usually grow very slowly until passages 4/5.

3b. Replace culture medium with 200 µl of 0.05% trypsin/1× EDTA and incubate 8 to 10 min in a 37◦ C, 5% CO2 incubator.

2B.2.8 Supplement 4

Current Protocols in Stem Cell Biology

4b. Collect the detached cells and transfer into a 15-ml tube. Add a drop of FBS to stop trypsin activity and fill the tube completely with DPBS. 5b. Centrifuge 5 min at 300 × g, 4◦ C. Resuspend the cell pellet in 2 ml high-glucose DMEM/20% FBS/1% PS. 6b. Replate the contents of one well into one uncoated well of a 12-well culture plate in 2 ml high-glucose DMEM/20% FBS/1% PS. Grow until confluent (passage 1) in a 37◦ C, 5% CO2 cell incubator. 7b. Repeat the procedure (steps 3b to 5b) and replate the cells from one well into two uncoated wells from a 6-well culture dish (passage 2) in 3 ml per well high-glucose DMEM/20% FBS/1% PS. Grow until confluent in a 37◦ C, 5% CO2 cell incubator. 8b. Repeat the procedure (steps 3b to 5b) and replate the cells from both wells of the 6well plate into one uncoated 25-cm2 culture flask (passage 3) in 10 ml high-glucose DMEM/20% FBS/1% PS. Grow until confluent in a 37◦ C, 5% CO2 cell incubator. 9b. Repeat the procedure (steps 3b to 5b) and replate the cells from one 25-cm2 culture flask into one uncoated 75-cm2 culture flask (passage 4) in 20 ml high-glucose DMEM/20% FBS/1% PS. Grow until confluent in a 37◦ C, 5% CO2 cell incubator. For this and subsequent steps, freeze cells at each passage. Freeze between 5 × 105 and 3 × 106 cells in a cryogenic vial containing 900 µl of FBS and 100 µl of DMSO. Freeze vials at −80◦ C, then transfer into liquid nitrogen.

10b. Repeat the procedure (steps 3b to 5b) and replate the cells from one 75-cm2 culture flask into three uncoated 75-cm2 culture flasks (passage 5) in 20 ml per flask highglucose DMEM/20% FBS/1% PS. Grow until confluent in a 37◦ C, 5% CO2 cell incubator. 11b. From now on until 16 weeks, passage cells 1:6, following steps 3b to 5b. 12b. From week 17, passage cells 1:3. At these later stages, cell growth is slower and fewer cells are obtained at a given interval.

RT-PCR ANALYSIS OF SORTED CELL SUBSETS All cell subsets sorted as described should be assayed for progenitor cell potential in culture and in vivo. To guarantee the absence of contaminating cells, 103 to 105 of each selected cell population should be set apart for RT-PCR analysis. It is particularly important to document the absence of cross-contamination between endothelial cells and pericytes/perivascular cells, since these two cell subsets are intimately associated with each other within blood vessel walls.

SUPPORT PROTOCOL 2

Materials Freshly sorted cells pelleted by microcentrifuging 1 min at 10,000 rpm, 4◦ C Absolutely RNA nanoprep kit (Stratagene) SuperScript II reverse transcriptase kit (Invitrogen) Taq DNA polymerase (Invitrogen) DNA oligonucleotides (Integraded DNA Technologies; see Table 2B.2.3 for size and sequence) 1% agarose gels Microcentrifuge (Sorvall Biofuge) PCR machine (thermal cycler model TC-312, Techne) 0.5-ml thin-walled PCR reaction tubes (GeneAmp, Applied Biosystems) Agarose gel electrophoresis system (DNA plus, USA Scientific) Gel documentation system (Gel Doc 2000, Bio-Rad)

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Table 2B.2.3 Primers for PCR Analysis of Myoendothelial Cells and Pericyte Cells

Access number

Product size

AGCCGAATGTGTAAAGGACAG 1172–1591

NM001025109

419

TGGAGACTCCTTCCAGCTTCA

GCTTCCACCACGATCTCATAC

U84722

274

CD56

GTATTTGCCTATCCCAGTGCC

CATACTTCTTCACCCACTGCTC 542–873

BC014205

331

Myf5

CACCTCCAACTGCTCTGATGG

GTGAATCGGTGCTGGCAACT

519–757

NM005593

238

Pax7

ACCAGGAGACCGGGTCCATC

CCCGAACTTGATTCTGAGC

868–1088

NM 002584

220

CD45

CATGTACTGCTCCTGATAAGAC

GCCTACACTTGACATGCATAC

940–1579

Y00638

639

CD146

AAGGCAACCTCAGCCATGTCG

CTCGACTCCACAGTCTGGGAC 168–603

M28882

435

NG2

GCTTTGACCCTGACTATGTTGGC TCCAGAGTAGAGCTGCAGCA

141–336

NM001897

195

β-actin

CCTCGCCTTTGCCGATCC

25–229

NM001101

204

Gene

Sense primera

Anti-sense primera

CD34

CATCACTGGCTATTTCCTGATG

CD144

Amplicon position

GGAATCCTTCTGACCCATGC

511–785

a All primers are listed from 5 to 3 .

1. Extract total RNA from 103 to 105 freshly sorted cells using the Absolutely RNA nanoprep kit according to the manufacturer’s instructions. Because of the limited numbers of sorted cells, this RNA kit is recommended for total RNA isolation.

2. Synthesize cDNA using the SuperScript II reverse transcriptase kit according to the manufacturer’s instructions. 3. Perform PCR with Taq DNA polymerase according to the manufacturer’s instructions. Run the following program on a thermal cyler:

1 cycle: 35 cycles:

1 cycle:

5 min 30 sec 30 sec 1 min 7 min

94◦ C (denature) 94◦ C 58◦ C (anneal) 72◦ C 72◦ C (extension)

The primers used for PCR are listed in Table 2B.2.3. Each set of oligonucleotides is designed to span two different exons so that genomic DNA contamination is of no concern.

4. Electrophorese PCR products on 1% agarose gels and photograph gels.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Proliferation medium

Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

1× Dulbecco’s modified Eagle medium (high-glucose DMEM, Invitrogen) 20% fetal bovine serum (FBS, Invitrogen) 5% horse serum (HS, Invitrogen) 2% chick embryo extract (Invitrogen) Store complete medium up to 1 month at 4◦ C

2B.2.10 Supplement 4

Current Protocols in Stem Cell Biology

COMMENTARY Background Information Besides early embryonic multipotent stem cells and adult-type tissue-committed progenitors, multilineage stem cells diversely designated as mesenchymal stem cells or marrow stroma cells (MSC), multipotent adult progenitor cells (MAPC), or muscle-derived stem cells (MDSC) have been detected in numerous adult tissues. MSC can differentiate into cells of mesodermal lineages (Beresford et al., 1992; Prockop, 1997; Pittenger et al., 1999; Toma et al., 2002). A rare subset of MAPC, initially identified in adult bone marrow, can give rise to derivatives of all three germ layers (Jiang et al., 2002). Cells resembling MSC and MAPC are present in mouse brain (Jiang et al., 2002), pancreas (Seaberg et al., 2004), dermis (Toma et al., 2001), and skeletal muscle (Qu-Petersen et al., 2002), as well as in human skin (Shih et al., 2005) and fat tissue (Zuk et al., 2001). Similarly, MDSC have been extracted from adult skeletal muscle using the “pre-plate” culture technique. MDSC regenerate skeletal muscle with a very high efficiency, but are also endowed with dramatic multilineage differentiation potential (Qu-Petersen et al., 2002). These multilineage cells have all been isolated retrospectively from cultured tissues. Therefore, the identity and localization of these stem cells within organs are unknown. Related stem cells have been, however, derived from avian and mouse embryonic and postnatal blood vessel walls and named mesoangioblasts (Cossu and Bianco, 2003). It was hypothesized that blood vessels, through their ubiquitous distribution in the organism, could act as stem cell carriers and disseminate regenerative potential. Recently, it has been demonstrated that definitive hematopoietic stem cells emerge in the early human embryo from a population of blood-forming endothelial cells, and these observations have been confirmed in the model of human embryonic stem cells (Tavian et al., 1999; Oberlin et al., 2002; Zambidis et al., 2005, 2006). Using the techniques described in this unit, the existence of blood vessel–associated multilineage progenitors in human embryonic, fetal, and adult tissues has been explored. Both human endothelial cells and vascular pericytes are endowed with the potential to generate various mesodermal derivatives, primarily skeletal muscle (Crisan et al. and Yap et al., unpub. observ.). In addition, a novel subset of muscle-derived myoendothelial cells that express markers of both myogenic and endothelial cell lineages

exhibit the strongest myogenic potential and may represent a developmental intermediate between both lineages (Zheng et al., 2007). A normal role of vascular cells in the development and/or regeneration of human muscle remains to be demonstrated (Gavina et al., unpub. observ.), but a similar myogenic potential is present within pericytes and endothelial cells purified from pancreas, fat, and other tissues (Crisan et al., unpub. observ.). In addition to myogenesis, there has been detected in pericytes and myoendothelial cells purified from human tissues a broader ability to differentiate into bone, cartilage, and adipocytes (Zheng et al., 2007 and Crisan et al., unpub. observ.), in agreement with published preliminary results (Farrington-Rock et al., 2004; Alliot-Licht et al., 2005; Collett and Canfield, 2005). This suggests the dissemination throughout organs of multilineage stem cells, which may be at the origin of MSC, MAPC, MDSC, and other adult stem cells identified retrospectively in culture. Blood vessel walls thus may harbor a dormant reserve of multilineage stem cells that could be recruited in emergency situations when professional tissue-specific progenitors have been exhausted. The blood vessel–derived novel myogenic progenitors that have been described can be sorted, by flow cytometry, from muscle and even more accessible sources such as fat tissue and bone marrow; these cells can then be cultured extensively, with no significant loss of developmental potential. The transplantation of autologous blood vessel-related progenitors could, therefore, be envisioned as a therapy for human muscle diseases (P´eault et al., 2007).

Critical Parameters It is important that all experiments be performed on fresh, viable tissues. Dead or dying cells should be kept as rare as possible, although these can be eliminated on the FACS following propidium iodide or 7-AAD staining. Most tissues stored in appropriate conditions can still be used for dissociation from 2 to 5 days post-mortem or autopsy. It is essential that cells be stained immediately after dissociation, otherwise many cells in the suspension will clump. If cells are dissociated in the evening and stored overnight at 4◦ C prior to antibody staining on the next morning, all pericytes are irremediably lost. The numbers of cells available for cell sorting vary vastly according to the tissue itself

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and developmental stage thereof. The cell subsets described in this article are rare and often sorted in small numbers (104 to 105 cells). The assays and culture conditions used are, however, sensitive enough to make such small cell populations usable. Typically, start from 3–5 × 106 and 2.5–3 × 107 total cells to sort cells from fetal and adult tissues, respectively. After the fifth passage in long-term culture, no less than 5 × 105 pericytes should be seeded per 75-cm2 flask. At lower cell densities, cells change morphology, grow slowly and eventually die. Conversely, it is not recommended to seed much more than 5 × 105 pericytes per 75-cm2 flask, as cells may grow too fast after 2 or 3 days and appear like fibroblasts, i.e., exhibit longer, thinner processes. It is therefore important, at each passage, to count the detached cells prior to passaging. The phenotype of cultured pericytes should also be regularly checked by immunocytochemistry and flow cytometry.

Troubleshooting The yield of total RNA prepared for RTPCR analysis is often low because of the limited numbers of sorted cells available which, moreover, can be made more fragile due to the pressure they are exposed to during the sorting process. It is, therefore, strongly recommended to use the Absolutely RNA nanoprep kit (Stratagene) for total RNA isolation. Performing RNA isolation immediately after sorting will also improve RNA yield.

Anticipated Results

Purification and Culture of Human Blood Vessel-Associated Progenitor Cells

Myoendothelial cells can be identified and sorted free of genuine endothelial cells and myogenic cells from the human fetal and adult skeletal muscle. Myoendothelial cells exhibit outstanding myogenic potential—about ten times higher than regular myogenic cells— both in culture and following injection into the cardiotoxin-treated SCID mouse skeletal muscle. This myogenic potential is sustained upon long-term culture of myoendothelial cells (Zheng et al., 2007). Pericytes are associated with capillaries and microvessels and are therefore present in all vascularized tissues. In the authors’ hands, pericytes have been sorted to homogeneity, as described above, from skeletal muscle, pancreas, placenta, brain, and adipose tissue. Both freshly isolated and long-term cultured pericytes exhibit the potential to differentiate into mesodermal tissues: myofibers,

cardiomyocytes, adipocytes, cartilage, and bone cells (Crisan et al., unpub. observ.).

Time Considerations On average and in the absence of significant trouble, a cell sorting experiment should require the following amounts of time. Dissociation of human tissues into single-cell suspensions takes 2 to 3 hr. Antibody staining takes 1 hr and cell sorting takes 1 to 3 hr (the rarer the population to be sorted, the longer the sort). RT-PCR analysis of sorted cell subsets requires 1 hr for total RNA isolation, 20 min for the preparation for reverse transcriptase reactions, 20 min for the preparation for PCR reactions, and 30 min for gel electrophoresis and gel documentation.

Literature Cited Alliot-Licht, B., Bluteau, G., Magne, D., LopexCazaux, S., Lieubeau, B., Daculs, G., and Guicheux, J. 2005. Dexamethasone stimulates differentiation of odontoblast-like cells in human dental pulp cultures. Cell Tissue Sep. 321:391-400. Beresford, J.N., Bennett, J.H., Devlin, C., Leboy, P.S., and Owen, M.E. 1992. Evidence for an inverse relationship between the differentiation of adipocytic and osteogenic cells in rat marrow stromal cell cultures. J. Cell Sci. 102:341-351. Collett, G.D. and Canfield, A.E. 2005. Angiogenesis and pericytes in the initiation of ectopic calcification. Circ. Res. 96:930-938. Cossu, G. and Bianco, P. 2003. Mesoangioblastsvascular progenitors for extravascular mesodermal tissues. Curr. Opin. Genet. Dev. 13:537542. Farrington-Rock, C., Crofts, N.J., Doherty, M.J., Ashton, B.A., Griffin-Jones, C., and Canfield, A.E. 2004. Chondrogenic and adipogenic potential of microvascular pericytes. Circulation 110:2226-2232. Jiang, Y., Vaessen, B., Lenvik, T., Blackstad, M., Reyes, M., and Verfaillie, C.M. 2002. Multipotent progenitor cells can be isolated from postnatal murine bone marrow, muscle, and brain. Exp. Hematol. 30:896-904. Oberlin, E., Tavian, M., Blazsek, I., and P´eault, B. 2002. Blood-forming potential of vascular endothelium in the human embryo. Development 129:4147-4157. P´eault, B., Rudnicki, M., Torrente, Y., Cossu, G., Tremblay, J., Partridge, T., Gussoni, E., Kunkel, L., and Huard, J. 2007. Stem and progenitor cells in skeletal muscle development, maintenance and therapy. Mol. Ther. 15:867-877. Pittenger, M.F., MacKay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., and Marshak, D.R. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284:143-147.

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Current Protocols in Stem Cell Biology

Prockop, D.J. 1997. Marrow stromal cells as stem cells for nonhematopoietic tissues. Science 276:71-74. Qu-Petersen, Z., Deasy, B.M., Jankowski, R., Ikezawa, M., Cummins, J., Pruchnic, R., Cao, B., Mytinger, J., Gates, C., Wernig, A., and Huard, J. 2002. Identification of a novel population of muscle stem cells in mice: Potential for muscle regeneration. J. Cell Biol. 157:851864. Seaberg, R.M., Smukler, S.R., Kieffer, T.J., Enikolopov, G., Asghar, Z., Wheeler, M.B., Korbutt, G., and Van Der Kooy, D. 2004. Clonal identification of multipotent precursors from adult mouse pancreas that generate neural and pancreatic lineages. Nat. Biotechnol. 22:11151124. Shih, D.T., Lee, D.C., Chen, S.C., Tsai, R.Y., Huang, C.T., Tsai, C.C., Shen, E.Y., and Chiu, W.T. 2005. Isolation and characterization of neurogenic mesenchymal stem cells in human scalp tissue. Stem Cells 23:1012-1020. Tavian, M., Hallais, M.F., and P´eault, B. 1999. Emergence of intraembryonic hematopoietic precursors in the pre-liver human embryo. Development 126:793-803. Toma, J.G., Akhavan, M., Fernandes, K.J., Barnabe-Heider, F., Sadikot, A., Kaplan, D.R., and Miller, F.D. 2001. Isolation of multipotent adult stem cells from the dermis mammalian skin. Nat. Cell Biol. 3:778-784.

Toma, C., Pittenger, M.F., Cahill, K.S., Byrne, B.J., and Kessler, P.D. 2002. Human mesenchymal stem cells differentiate to a cardiomyocyte phenotype in the adult murine heart. Circulation 105:93-98. Zambidis, E.T., P´eault, B., Park, T.S., Bunz, F., and Civin, C.I. 2005. Hematopoietic differentiation of human embryonic stem cells progresses through sequential hemato-endothelial, primitive, and definitive stages resembling human yolk sac development. Blood 106:860-870. Zambidis, E., Oberlin, E., Tavian, M., and Peault, B. 2006. Blood-forming endothelium in human ontogeny: Lessons from in utero development and embryonic stem cell culture. Trends Cardiovasc. Med. 16:95-101. Zheng, B., Cao, B., Crisan, M., Sun, B., Li, G., Logar, A., Yap, S., Pollet, J.B., Drowley, L., Cassino, T., Gharaibeh, B., Deasy, B.M., Huard, J., and P´eault, B. 2007. Prospective identification of myogenic endothelial cells in human skeletal muscle. Nat. Biotechnol. 25:10251034. Zuk, P.A., Zhu, M., Mizuno, H., Huang, J., Futrell, J.W., Katz, A.J., Benhaim, P., Lorenz, H.P., and Hedrick, M.H. 2001. Multilineage cells from human adipose tissue: Implications for cell-based therapies. Tissue Eng. 7:211-228.

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Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

UNIT 2B.3

Fernando Anjos-Afonso1 and Dominique Bonnet1 1

Haematopoietic Stem Cell Laboratory, Cancer Research UK, London Research Institute, London, United Kingdom

ABSTRACT This unit describes how to isolate and expand mesenchymal stromal cells (MSCs) from mouse bone marrow. For reasons that are not clear, it has been difficult to isolate these cells (also known as mesenchymal stem cells). Furthermore, different mouse strains seem to have specific requirements for successful extraction and culture of these cells. A general and easy protocol is presented here for isolating stromal cells from different inbred and transgenic mice commonly used in the stem cell biology field. Curr. Protoc. C 2008 by John Wiley & Sons, Inc. Stem Cell Biol. 7:2B.3.1-2B.3.11.  Keywords: mouse mesenchymal stromal cells r isolation r expansion r differentiation

INTRODUCTION Mesenchymal stromal cells (MSCs) from human and rat bone marrow have been extensively described, in part because they are relatively easy to isolate by their adherence to plastic and can be extensively expanded in culture (Pittenger et al., 1999; Colter et al., 2000, 2001; Javazon et al., 2001). Protocols for their isolation and expansion are fairly similar among different laboratories. In contrast, detailed literature on this subject for mouse cells is limited, and the methods differ among different groups. Most researchers use undefined mouse bone marrow adherent fractions as a source of stromal cells. However, these adherent fractions, especially from early cell passages, contain many persistent hematopoietic contaminants. On top of that, isolation and expansion of stromal cells from different mouse strains seem to require different basal medium, type of serum, culturing time, and cell plating densities (Phinney et al., 1999; Peister et al., 2004). All of these peculiarities render interpretation of some of the published results difficult and have greatly limited the ability to test these cells in different genetic models. The Basic Protocol below describes one common and simple sorting method that can be used to isolate these mesenchymal stromal cells from C57Bl/6J, 129, FVB/N, C57Bl/6Actb-eGFP, Rosa26-eGFP (FVB/N background), Rosa26-LacZ (129/Bl6 background), and NOD/SCID mouse strains. Support Protocols 1 and 2 describe staining procedures for differentiated cell types. NOTE: Reagents can be purchased from companies other than those suggested in the materials lists in the protocols. NOTE: Use electric or manual pipettors (for pipetting up to 25 ml) and 2- to 1000-μl micropipettors and pipet tips with sterile aerosol barriers. NOTE: The following procedures must be performed in a tissue culture hood. NOTE: All solutions and equipment must be sterile and proper aseptic techniques should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator. Somatic Stem Cells Current Protocols in Stem Cell Biology 2B.3.1-2B.3.11 Published online October 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02b03s7 C 2008 John Wiley & Sons, Inc. Copyright 

2B.3.1 Supplement 7

NOTE: Culture medium should be warmed to 37◦ C before use. Warm only the necessary volume to avoid multiple warming/cooling cycles from the same bottle of medium. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. BASIC PROTOCOL

Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

This protocol describes a general method for isolation, culture, and differentiation of mesenchymal stromal cells from the bone marrow of a variety of mouse strains used in stem cell biology. Some parts of this method have been adapted from work of Professor D. Prockop’s team (Phinney et al., 1999; Peister et al., 2004) and modified from the authors’ early publications (Anjos-Afonso et al., 2004) to create a more general protocol that covers the isolation of MSCs from different mouse strains. Nevertheless, each researcher should adjust this method according to growth behavior of the cells under study (e.g., growth kinetics, differentiation potential; see Critical Parameters and Troubleshooting). Cells derived from the same mouse strain can sometimes vary from one laboratory to another.

Materials 6- to 10-week-old mice of appropriate strain(s) CMF-PBS/2% FBS: phosphate-buffered saline, without Ca2+ and Mg2+ (CMF-PBS; GIBCO-Invitrogen, cat. no. 20012) containing 2% (v/v) heat-inactivated fetal bovine serum (FBS; StemCell Technologies, cat. no. 06471) Complete MSC expansion medium (see recipe) 0.4% (w/v) trypan blue (Sigma, cat. no. T6140) solution in CMF-PBS (GIBCO-Invitrogen, cat. no. 20012) Phosphate-buffered saline without Ca2+ and Mg2+ (CMF-PBS) 0.25% (w/v) trypsin/1 mM EDTA solution (ethylenediaminetetraacetic acid in Hanks’ balanced salt solution; GIBCO-Invitrogen, cat. no. 25200) Fetal bovine serum (FBS; StemCell Technologies, cat. no. 06471): low endotoxin; batch preselected for MSC growth; do not heat inactivate the serum; store at −20◦ C and thaw before use overnight at 4◦ C (do not thaw at 37◦ C) Fluorphore-conjugated antibodies (BD Bioscience Pharmingen; choice of fluorophore at the discretion of the researcher): Anti-mouse CD45 (clone 30-F11) Anti-mouse CD11b (clone M1/70) Anti-mouse CD31 (clone 390) Anti-mouse CD105 (clone MJ7/18) Anti-mouse CD73 (clone TY/23) Anti-mouse Sca1 (clone E13-161.7) Anti-mouse CD44 (clone IM7) Rat-IgG2a (clone R35-95) isotype control Rat-IgG2b (clone A95-1) isotype controls Live/dead discriminative dye stock solution, e.g.: 200 mg/ml of 4 ,6-diamidino-2-phenylindole (DAPI; Invitrogen, cat. no. D1306) 40 mg/ml of 7-amino-actinomycin D (7-AAD; Invitrogen, cat. no. A1310) Adipogenic induction medium (see recipe), 37◦ C Osteogenic induction medium (see recipe), 37◦ C Alkaline phosphatase detection kit (Sigma, Kit 86) Chondrogenic induction medium (see recipe), 37◦ C Sterile dissecting scissors Insulin syringe or 1-ml syringe with a 27- to 29-G needle 5- and 15-ml polyprene tubes Refrigerated centrifuge, 4◦ C

2B.3.2 Supplement 7

Current Protocols in Stem Cell Biology

25-cm2 tissue culture–treated culture flasks/dishes (or other appropriate size) Humidified cell culture incubator (37◦ C, 5% CO2 ) 30◦ C and 37◦ C water bath 0.22-μm filter Cell sorter (e.g., MoFlow, Dako-Cytomation) Additional reagents and equipment for assessing cell number and viability (UNIT 1C.3) Isolate mouse bone marrow stromal cells 1. Euthanize the mice (by cervical dislocation or other appropriate means) and collect the femurs, tibias, and iliac crests from 6- to 10-week-old mice of the desired strain(s), using proper aseptic techniques. Remove all connective tissue. Do not use mice that are older than 12 weeks.

2. With sterile scissors, slightly cut open both epiphysis areas of each bone, flush the marrow a few times with an insulin syringe (or a 1-ml syringe coupled with a 27- to 29-G needle) containing CMF-PBS/2% FBS, and transfer to a 5-ml polyprene tube. One ml of CMF-PBS/2% FBS is enough volume to flush off the marrow of the six bones of one mouse.

3. Centrifuge the cells from the marrow 5 min at 380 × g, 4◦ C. 4. Discard the supernatant, resuspend the cells with 1 ml complete MSC expansion medium and count an aliquot of the cells with 0.4% trypan blue solution to assess cell number and viability (UNIT 1C.3). 5. Seed 1 × 106 cells/cm2 onto a 25-cm2 tissue culture–treated culture flask or dish and incubate the cells at 37 ◦ C, 5% CO2 . Normally, marrow cells collected from one immunodeficient mouse (NOD/SCID) are enough to seed one 25-cm2 tissue flask; two flasks can be seeded with marrow cells collected from one mouse for other (nonimmunodeficient) mouse strains. The expected yield for one immunodeficient mouse is ∼25 × 106 cells and ∼50 × 106 cells for other strains.

Grow the primary culture 6. Add 5 ml of complete expansion medium for a 25-cm2 tissue culture flask. Adjust the amount of medium proportionally to the area of the flask/dish used.

7. Eliminate nonadherent cells by changing the entire medium at day 3 and then replacing with fresh complete expansion medium every 4 days for 10 to 15 days until cells reach 70% confluency. These are the primary cultures (passage 0; P0).

Passage the cells 8. Remove the medium and wash the adherent cells once with 3 to 5 ml CMF-PBS. 9. Incubate the cells with 2 to 3 ml of the 0.25% trypsin/EDTA solution for 5 min at 37◦ C, until the cells detach from the flasks. Discard cells that do not detach. 10. Neutralize the trypsin by adding 1 ml FBS, then collect and centrifuge the cells 5 min at 380 × g, 4◦ C. 11. Split the cells at ratio of 1:3 and reseed them onto new flasks (passage 1; P1). 12. Culture P1 cells for 2 to 3 weeks until they reach 70% confluency. Change to fresh complete expansion medium every 4 days.

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Purify bone marrow stromal cells 13. Collect the cells from P1 by trypsinization, as described in steps 8 to 10. 14. Also collect the P1 medium and filter it through a 0.22-μm filter. Store at −20◦ C until use in step 22. 15. Resuspend the cells with CMF-PBS/2% FBS and count an aliquot (UNIT 1C.3). Adjust the cell density to 1 × 106 cells/50 μl with CMF-PBS/2% FBS and add 50 μl to a 5-ml polyprene tube. 16. Add the anti-mouse CD45 and anti-mouse CD11b antibodies each at a concentration of 2 μg/ml (e.g., add 0.25 to 1 μl of each antibody to the 50 μl of cell suspension, depending on the concentration of each stock antibody). Incubate 30 min at 4◦ C. Do not use FITC-conjugated antibodies for MSCs derived from eGFP mouse strains because GFP and FITC have similar emission wavelengths.

17. To wash the cells, add 3 ml CMF-PBS/2% FBS to the polyprene tube. Centrifuge 5 min at 380 × g, 4◦ C. 18. Discard the supernatant and resuspend the cells with a desirable volume of CMFPBS/2% FBS containing a live/dead discriminative dye (e.g., DAPI stock solution diluted 1:1000 or 7-AAD stock solution diluted 1:10). Consult the technical staff from the flow sorting laboratory for the desirable cell concentration for cell-sorting.

19. Sort for the live cells that are negative for both CD45 and CD11b expression (see example in Fig. 2B.3.1A). Use sorts carried out at the speed of 3000 to 5000 cells/sec with a 100-μm nozzle set-up. 20. After collecting the sorted cells, centrifuge them 5 min at 380 × g, 4◦ C. 21. Discard the supernatant, resuspend the cells in complete expansion medium, and count an aliquot (UNIT 1C.3). 22. Reseed the cells at 5000 cells/cm2 with a 1:1 mix of filtered P1-derived medium (collected in step 14) and complete MSC expansion medium (passage 2, P2). 23. Change to fresh complete expansion medium at day 3. Mouse bone marrow stromal cells seem to require some secreted factors from the hematopoietic contaminants to survive ex vivo in early cell passages. Culturing P2 cells with some medium from P1 and in high density at this point will assure better survival of the cells after being exposed to the stressful sorting conditions and growing for the first time without the presence of the hematopoietic cells.

24. Incubate the cells 5 to 7 days at 37◦ C, 5% CO2 , until the cells reach 70% confluency (see ∼50% confluent P2 culture in Fig. 2B.3.1B). Cells can also be purified using antibody-based magnetic column separation techniques instead of sorting. If this last method is chosen, follow the guidelines of the manufacture from which the separation reagents were purchased and then follow steps 20 to 24.

Expand bone marrow stromal cells 25. Expand the cells from P2 onwards, seeding them at 50 to 1000 cells/cm2 in complete expansion medium. Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

Higher-fold expansion is achieved when cells are plated at low cell density (e.g., 50 to 100 cell/cm2 ). However, plating at low cell density requires more flasks/dishes, reagent, and space. Thus, each laboratory should adjust according to their needs.

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Figure 2B.3.1 (A) Representative flow cytometric analysis plot of a P1 culture derived from C57Bl/6 bone marrow cells. Gated area (negative for both CD45 and CD11b expressions) should be used to sort out mouse MSCs from the hematopoietic contaminants. (B) A phase-contrast picture taken from C57Bl/6 derived MSCs at P2 (∼50% confluent). (C) Flow cytometric histograms showing the lack of expression of CD45, CD11b, and CD31 antigens on the surface of MSCs derived from Rosa26-LacZ mice. On the other hand, these cells are positive for CD105, CD44, CD73, and Sca1 expressions. Gray and white areas are the isotype control and specific antibody staining, respectively. In vitro differentiation of NOD/SCID-derived MSCs into adipogenic (D), osteogenic (E) and chondrogenic (F) lineages respectively. Magnifications: 100× (B, D, E); 400× (F).

26. Change to fresh complete expansion medium every 4 days until the cells reach 70% confluency. 27. Passage the cells as described in steps 8 to 10 and reseed the cells at the same cell density as chosen for P3. 28. Continue culture in the same manner.

Phenotype the cells 29. Collect cells from P3 by trypsinization as described in steps 8 to 10. 30. Stain the cells for CD45, CD11b, CD31, CD105, Sca1, CD44, and CD73 expressions. Use 0.2–1 × 106 cells for each antibody staining. In addition, allocate some

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cells for appropriate fluorochrome-matched isotype control staining (see example in Fig. 2B.3.1C) and analyze all by flow cytometry. MSCs are negative for CD45, CD11b, and CD31 and positive for CD105, Sca-1, CD44, and CD73 expressions, respectively. The phenotype of the cells for these antigens is stable until passages 8 to 10. Do not use FITC-conjugated antibodies for MSCs derived from eGFP mouse strains because GFP and FITC have similar emission wavelengths.

Differentiate bone marrow stromal cells For adipogenic and osteogenic differentiations 31a. Plate the cells at 2500 cell/cm2 in MSC expansion medium. Seed at least three wells for each differentiation condition. Incubate 36 to 48 hr. To achieve good differentiation, use cells up to passage 8, as cells tend to lose their differentiation capacity during further passaging.

32a. Change the medium to adipogenic or osteogenic induction medium, respectively. For a well of 10-cm2 (in a 6-well plate), add 2.5 ml of induction medium/well and for wells of smaller surface area adjust the volume accordingly.

33a. Replace all of the medium with the appropriate fresh induction medium every 3 days for 2 weeks. 34a. Evaluate cultures for adipogenic differentiation (Oil Red O staining; Support Protocol 1) and/or osteogenic differentiation (alkaline phosphatase staining; Sigma kit 86). Alkaline phosphatase staining is red and the nuclei are blue when hematoxylin is used (see example in Fig. 2B.3.1E).

For chondrogenic differentiation 31b. Place 2.5–5 × 105 cells in a 15-ml polyprene tube with 500 μl of complete chondrogenic induction medium. Centrifuge 5 min at 380 × g, 4◦ C, to pellet the cells. 32b. Put the tube in a rack and place it vertically in the incubator. 33b. Replace the medium with 1 ml of fresh complete chondrogenic induction medium every 3 days for 3 weeks. 34b. Evaluate the cultures for chondrogenic differentiation (Safranin O staining; Support Protocol 2). SUPPORT PROTOCOL 1

OIL RED O STAINING Oil red O staining is used to detect the lipid in adipocytes.

Materials

Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

Mouse MSC cells cultured for adipogenic differentiation (Basic Protocol, step 34a) Phosphate-buffered saline, without Ca2+ and Mg2+ (CMF-PBS; GIBCO-Invitrogen, cat. no. 20012) Oil Red O stain working solution (see recipe) 10% (v/v) neutral buffered formalin (NBF; see recipe) 60% (v/v) isopropanol Gill’s hematoxylin solution (Vector Laboratories, cat. no. H-3401) Phase-contrast microscope (100× to 400× magnification recommended)

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Fix and stain the cultures 1. At the end of 2 weeks of culture under adipogenic differentiation conditions, aspirate the medium from the well. 2. Gently rinse each well with 2 ml CMF-PBS by dispensing it along the sides of each well of the plate. 3. Aspirate the CMF-PBS, add 2 ml NBF, and incubate 30 to 60 min at room temperature. 4. Prepare the Oil Red O stain working solution (see recipe). 5. Remove the formalin solution from the cells and gently rinse once with 2 ml distilled water. Then add 2 ml of 60% isopropanol and incubate 5 min at room temperature. 6. Pour off the isopropanol and pipet 2 ml Oil Red O working solution along the side of each well. 7. Slowly rotate the plate to spread the stain and incubate 15 min at room temperature. 8. Rinse with tap water until the water runs clear.

Counterstain with hematoxylin (optional) 9. Add 2 ml Gill’s hematoxylin solution and incubate 1 min at room temperature. 10. Pour the hematoxylin off and rinse the plate with tap water until the water runs clear. Counterstaining with hematoxylin may be used to give better contrast, if desired.

Visualize the results 11. Keep the plates wet with water or CMF-PBS until they are ready to be viewed. 12. View the plate on a phase-contrast microscope (magnification of 100× to 400× recommended). Lipids appear in red and the nuclei are blue when hematoxylin is used (see example in Fig. 2B.3.1D).

SAFRANIN O STAINING Safranin O staining, counterstained with Fast Green FCF, is used to detect chondrocytes.

SUPPORT PROTOCOL 2

Materials Mouse MSC cells cultured for chondrogenic differentiation (Basic Protocol, step 34b) 10% (v/v) neutral buffered formalin (NBF; see recipe) 50%, 70%, 75%, 95%, 100% (v/v) ethanol Xylene Weigert’s iron hematoxylin solution kit (Sigma, cat. no. HT1079), containing stock solutions A and B Fast Green staining solution (see recipe) 1% (v/v) acetic acid: prepare by diluting glacial acetic acid (Sigma, cat. no. A9967) 0.1% (w/v) Safranin O (Sigma, cat. no. 84120) staining solution: filter through filter paper before use Resinous mounting medium (Vector Laboratories, cat. no H-5000) Light microscope (100× to 400× magnification recommended) Additional reagents and equipment for embedding and sectioning cells (supplied by histology service)

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Prepare slides 1. At the end of 3 weeks of culture under chondrogenic differentiation conditions, gently aspirate the induction medium from the cells, and add 1 ml NBF into the tube. 2. Aspirate the buffered formalin, replace it again with 1 ml NBF, and fix the pellet for 24 hr at room temperature. 3. Transfer the fixed pellet to 70% ethanol until is embedded in paraffin. 4. Take the pellet to the histology service or laboratory for embedding in paraffin and sectioning. Use 6-μm sections for Safranin O staining. 5. Deparaffinize and hydrate by immersing the slides into the following:

Xylene, twice for 10 min each time 100% ethanol, twice for 1 min each time 95% ethanol, 2 min 75% ethanol, 2 min 50% ethanol, 2 min Distilled water, twice for 2 min each time. Counterstain slides 6. Stain with Weigert’s iron hematoxylin working solution (equal parts of stock solutions A and B) for 10 min. 7. Wash in running tap water for 10 min. 8. Stain with Fast Green staining solution for 5 min. 9. Rinse quickly with 1% acetic acid solution for 10 sec.

Stain with Safranin O 10. Stain in 0.1% Safranin O staining solution for 5 min. 11. Dehydrate and clear with:

95% ethanol, twice for 2 min each 100% ethanol, twice for 2 min each Xylene, twice for 2 min each. 12. Mount the slides with resinous mounting medium. 13. Examine the sections with a microscope (magnification of 100× to 400× recommended). Cartilage and mucin appears in orange/red, cytoplasm is dark green, and the nuclei are black (see example in Fig 2B.3.1F).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. Sterilize all stock reagents (except those made in methanol) by passing through a 0.22-μm filter before use. Store stock reagents in aliquots up to 3 months at 4◦ C or for longer periods at −20◦ C, unless otherwise indicated.

Adipogenic induction medium Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

Dulbecco’s modified Eagle medium (DMEM), low-glucose formulation 2% (v/v) FBS (FBS; StemCell Technologies, cat. no. 06471): low endotoxin; batch preselected for MSC growth; do not heat inactivate the serum; store at −20◦ C and thaw before use overnight at 4◦ C (do not thaw at 37◦ C) continued

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1× pen/strep (final concentrations of 100 U/ml and 100 μg/ml, respectively; 10,000 U/ml penicillin/10,000 μg/ml streptomycin; GIBCO-Invitrogen, cat. no. 15140) 1 μM dexamethasone: use 1 mM dexamethasone (Sigma, cat. no. G9891) stock solution 50 μM indomethacin: use 40 mM indomethacin (Sigma, cat. no. 17378) stock solution in methanol 500 nM IBMX: use 10 mM isobutylmethylxanthine (IBMX: Sigma, cat. no. 17018) stock solution in methanol 5 μg/ml insulin: use 10 mg/ml insulin stock solution (Sigma, cat. no. 19278) Prepare fresh Alternatively, make larger volumes than needed for 1 day and store the medium up to 1 month at 4◦ C.

Chondrogenic induction medium Serum-free Dulbecco’s modified Eagle medium (DMEM), high-glucose formulation 6.25 μg/ml insulin: use 10 mg/ml insulin stock solution 6.25 μg/ml of transferrin: use 6.25 mg/ml transferrin (Sigma cat. no. T8158) stock solution; store up to 7 days at 4◦ C 6.25 μg/ml of sodium selenite: use 6.25 mg/ml sodium selenite (Sigma, cat. no. S5261) stock solution; store stock solution up to 1 month at 4◦ C 5.33 μg/ml of linoleic acid: use 0.533 mg/ml linoleic acid (Sigma, cat. no. L5900) stock solution 1.25 mg/ml of bovine serum albumin (BSA): use 125 mg/ml albumin bovine serum (Sigma, cat. no. A9647) stock solution 100 nM dexamethasone: use 1 mM dexamethasone (Sigma, cat. no. G9891) stock solution 50 μM ascorbic acid: use 200 mM ascorbic acid (L-ascorbic acid-2-phosphate sesquimagnesium salt; Sigma, cat. no. A8960) stock solution; store aliquots of stock solution at −20◦ C 10 ng/ml of transforming growth factor (TGFβ3 ; Peprotech, cat. no. 100-36) Prepare fresh Complete MSC expansion medium Mix the whole 100 ml bottle of the Mouse Mesenchymal Stem Cell Stimulatory Supplements (Stem Cell Technologies, cat. no. 05502) with the entire 400 ml bottle of MesenCult Basal Medium (Stem Cell Technologies, cat. no. 05501) and supplement the mixture with 5 ml pen/strep (final concentration 100 U/ml and 100 μg/ml respectively; 10,000 U/ml penicillin/10,000 μg/ml streptomycin; GIBCO-Invitrogen, cat. no. 15140). Store the complete medium up to 1 month at 4◦ C.

Fast Green staining solution Make 0.001% (w/v) Fast Green FCF (Sigma, cat. no. F7258) stain solution and filter through with filter paper. Store at room temperature.

Neutral buffered formalin, 10% (pH 7) 3.5 g sodium phosphate, dibasic (Na2 HPO4 ; Sigma, cat. no. 57907) 3.5 g sodium phosphate, monobasic (NaH2 PO4 ; Sigma, cat. no. 58282) 100 ml 37% formaldehyde (Sigma, cat. no. F1635) 900 ml H2 O Somatic Stem Cells

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Oil Red O stain solutions Stock solution: Make 0.5% (w/v) Oil Red O (Sigma, cat. no. O0625) stock solution in isopropanol and filter the mixture with filter paper. Store up to 6 months at 4◦ C. Working solution: Mix 3 parts Oil Red O stock solution with 2 parts water and let the mixture stand for 10 min. Filter the solution using filter paper. Prepare fresh for each use.

Osteogenic induction medium DMEM, low-glucose formulation 2% (v/v) FBS (FBS; StemCell Technologies, cat no. 06471): low endotoxin; batch preselected for MSC growth; do not heat inactivate the serum; store at −20◦ C and thaw before use overnight at 4◦ C (do not thaw at 37◦ C) 1× pen/strep (final concentrations of 100 U/ml and 100 μg/ml, respectively; 10,000 U/ml penicillin/10,000 μg/ml streptomycin; GIBCO-Invitrogen, cat. no. 15140) 50 μM ascorbic acid: use 200 mM ascorbic acid (L-ascorbic acid-2-phosphate sesquimagnesium salt; Sigma, cat. no. A8960) stock solution; store aliquots of stock solution at −20◦ C 10 nM dexamethasone: use 1 mM stock solution 10 mM β-glycerolphosphate: use 0.5 M β-glycerolphosphate (Sigma, cat. no. G9891) stock solution Prepare fresh Alternatively, make larger volumes than needed for 1 day and store the medium up to 1 month at 4◦ C.

COMMENTARY Background Information

Isolation, Culture, and Differentiation Potential of Mouse Marrow Stromal Cells

This protocol is suitable for isolating and expanding stromal cells from different mouse strains. However, it is still recommended that each laboratory determine the optimal plating cell densities and time intervals for cell passaging. The growth properties of mouse bone marrow stromal cells not only are strain dependent, but also donor dependent. To avoid sample variation, it is also recommended to pool the flushed marrows from a few mice from the same experimental conditions, if possible. For reasons that are not clear, mouse stromal cells are prone to develop chromosomal abnormalities; thus, karyotype analysis is recommended. However, this characteristic does not seem to alter their in vitro differentiation potentials (F. Anjos-Afonso, pers. observ.). The level of expression of common antigens found on the cell surface of mouse MSCs (e.g., CD105, CD73, CD44, and Sca-1) varies from one mouse strain to another. Of note, mouse stromal cells from mouse strains listed in the unit introduction do express CD34 antigen (not found on human MSCs), but not on others strains (Peister et al., 2004). Unlike human MSCs, mouse MSC do not express CD90 (Thy-1) antigen.

Critical Parameters and Troubleshooting Of all the strains tested, cells from C57Bl/6J and C57Bl/6-Actb-eGFP were the most difficult to isolate. If cells do not survive to P2 after being reseeded at the specified densities, they should be reseeded at higher cell densities, e.g., 10,000 to 15,000 cell/cm2 . Alternatively, use cells from P2 instead from P1 for cell sorting. In this case, passage the cells once more (P1 to P2) at density of 5000 cell/cm2 and use P2 cells when they reach 70% confluency. Examine the cells regularly with a microscope. If the cells slow down or stop growing at any time and/or become flattened, discard the cultures. All procedures must be performed aseptically, as fungal contamination frequently occurs. If such contamination develops, discard the cultures. Usually the quality of FBS is critical in MSC culture. But by using the basic MSC medium with the supplements recommended for this protocol, screening different lots of FBS is now less crucial for the growth of the cells. Nonetheless, different batches of FBS might influence the differentiation potential of the cells and this should be verified.

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The cell density used to initiate adipogenic and osteogenic differentiations can be increased, and the duration of differentiation can be extended, according to the behavior of each culture. Cells can be seeded up to 5000 cell/cm2 before induction and the duration extended to 3 weeks. However, sometimes this can lead the cells to detach during the time course of the differentiation.

Anticipated Results This protocol generates large numbers of stromal cells that are suitable for gene marking experiments using viral-based vectors and for differentiation studies. It is expected that after cell sorting stromal cultures will be devoid of hematopoietic contaminants. Differentiation potential of the stromal cells also depends on the mouse strain from which they were derived. It is expected that cells from any strain could differentiate at least into two of the three conventional mesenchymal lineages (adipogenic and osteogenic). In general, cells derived from all the strains tested were able to differentiate well into osteoblasts. Cells from NOD/SCID and FVB/N are less prone to differentiate into adipocytes compared to other strains. It is generally difficult to form stable cartilaginous pellets with mouse stromal cells. Cells from C57Bl/6 or related strains usually don’t form cartilaginous tissue well in vitro, whereas cells from NOD/SCID do it efficiently.

Time Considerations

The whole process it takes ∼6 weeks from isolating bone marrow cells to P2. Subsequently, depending on the seeding density chosen, each passage can vary from 5 to 12 days (e.g., if cells are seeded at 5000 cells/cm2 it should take ∼5 days to reach confluency, whereas it can take up to 12 days when the

cells are seeded at 100 cells/cm2 ). It is recommended to passage cells when they reach ∼70% to 80% confluency and not let them overgrow over weekends. It takes 2 to 3 weeks to test the differentiation capacity of the cells.

Literature Cited Anjos-Afonso, F., Siapati, E.K., and Bonnet, D. 2004. In vivo contribution of murine mesenchymal stem cells into multiple cell-types under minimal damage conditions. J. Cell Sci. 117:5655-5664. Colter, D.C., Class, R., DiGirolamo, C.M., and Prockop, D.J. 2000. Rapid expansion of recycling stem cells in cultures of plastic-adherent cells from human bone marrow. Proc. Natl. Acad. Sci. U.S.A. 97:3213-3218. Colter, D.C., Sekiya, I., and Prockop, D.J. 2001. Identification of a subpopulation of rapidly self-renewing and multipotential adults stem cells in colonies of human marrow stromal cells. Proc. Natl. Acad. Sci. U.S.A. 98:78417845. Javazon, E.H., Colter, D.C., Schwarz, E.J., and Prockop, D.J. 2001. Rat marrow stromal cells are more sensitive to plating density and expand more rapidly from single-cell-derived colonies than human marrow stromal cells. Stem Cells 19:219-225. Peister, A., Mellad, J.A., Larson, B.L., Hall, B.M., Gibson, L.F., and Prockop, D.J. 2004. Adult stem cells from bone marrow (MSCs) isolated from different strains of inbred mice vary in surface epitopes, rates of proliferation, and differentiation potential. Blood 103:1662-1668. Phinney, D.G., Kopen, G., Isaacson, R.L., and Prockop, D.J. 1999. Plastic adherent stromal cells from the bone marrow of commonly used strains of inbred mice: Variations in yield, growth, and differentiation. J. Cell. Biochem. 72:570-585. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Graig, S., and Marshak, D.R. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284:143147.

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Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

UNIT 2C.1

Laura E. Mead,1, 2 Daniel Prater,1, 2 Mervin C. Yoder,1, 2, 3 and David A. Ingram1, 2, 3 1

Department of Pediatrics, Indiana University School of Medicine, Indianapolis, Indiana Herman B Wells Center for Pediatric Research, Indiana University School of Medicine, Indianapolis, Indiana 3 Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana 2

ABSTRACT Circulating endothelial progenitor cells (EPCs) in adult human peripheral blood were originally identified in 1997 by Asahara et al., which challenged the paradigm that vasculogenesis is a process restricted to embryonic development. Since their original identification, EPCs have been extensively studied as biomarkers to assess the risk of cardiovascular disease in human subjects and as a potential cell therapeutic for vascular regeneration. Endothelial colony-forming cells (ECFCs), which are a subtype of EPCs, were recently identified from circulating adult and human umbilical cord blood. In contrast to other types of EPCs, which display various monocyte/macrophage phenotypes and functions, ECFCs are characterized by robust proliferative potential, secondary and tertiary colony formation upon replating, and de novo blood vessel formation in vivo when transplanted into immunodeficient mice. In this unit, we describe detailed methodologies for isolation and characterization of ECFCs from both human peripheral and umbilical C 2008 by John Wiley & cord blood. Curr. Protoc. Stem Cell Biol. 6:2C.1.1-2C.1.27.  Sons, Inc. Keywords: endothelial progenitor cell (EPC) r adult progenitor cells r endothelial cell transplantation r endothelial colony-forming cells (ECFC)

INTRODUCTION Since 1997, postnatal vasculogenesis has been purported to be an important mechanism for angiogenesis via marrow-derived circulating endothelial progenitor cells (EPC; Asahara et al., 1997). Based on this paradigm, EPCs have been extensively studied as biomarkers of cardiovascular disease and as a cell-based therapy for repair of damaged blood vessels (Hill et al., 2003; Rafii and Lyden, 2003; Werner et al., 2005). Various methods exist for isolation and identification of EPCs. This unit describes a method for isolation and cultivation of endothelial colony-forming cells (ECFC), which are EPCs with high proliferative potential. Mononuclear cells (MNC) are isolated from umbilical cord blood (UCB; Basic Protocol 1) or peripheral blood (Alternate Protocol 1) and cultured in endothelial-specific growth conditions, yielding ECFC colonies within one to several weeks. The phenotype of endothelial outgrowth cells is confirmed by examination of endothelial-specific cell surface antigen expression (Basic Protocol 2) and assessment of endothelial cell (EC) function via acetylated low-density lipoprotein (AcLDL) ingestion (Alternate Protocol 2) and tube formation on Matrigel (Alternate Protocol 3). Further, verification of ECFC functionality can be achieved by assessing their ability to contribute to de novo vasculogenesis in an in vivo murine implant model (Basic Protocol 3). Somatic Stem Cells Current Protocols in Stem Cell Biology 2C.1.1-2C.1.27 Published online July 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02c01s6 C 2008 John Wiley & Sons, Inc. Copyright 

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NOTE: All procedures described in this unit are performed under sterile conditions in a Class II biohazard flow hood. All solutions and equipment that come into contact with live cells must be sterile and proper aseptic technique must be used. NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator, unless otherwise noted. NOTE: All media used for suspension, wash, or culture of cells is prewarmed to 37◦ C in a water bath. NOTE: This protocol uses human blood. It should be submitted, reviewed, and approved by the appropriate institutional review board. All samples should be obtained with informed consent of the donor. NOTE: Human blood is potentially infectious and standard precautions for the safe use of human tissue should be followed. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee and must conform to governmental regulations for the care and use of laboratory animals. BASIC PROTOCOL 1

ISOLATION, CLONING, AND PROPAGATION OF ENDOTHELIAL COLONY-FORMING CELLS FROM HUMAN UMBILICAL CORD BLOOD Primary culture of UCB MNCs results in the outgrowth of ECFC-derived colonies between day 5 and day 14. Individual endothelial cell colonies can be clonally isolated and serially subcultured. Cells are maintained in specific culture medium and have the potential to undergo up to 100 population doublings (Ingram et al., 2004).

Materials Anticoagulated UCB (e.g., the authors typically use 10 U heparin/ml blood) Phosphate-buffered saline (PBS), without calcium and magnesium, pH 7.2 Ficoll-Paque PLUS (Ficoll, Amersham Biosciences, cat. no. 17-1440-03) EBM-2 10:1 (see recipe; Lonza) 0.4% (w/v) trypan blue solution (Sigma, cat. no. T8154) cEGM-2 (see recipe; Lonza) 15-ml conical centrifuge tubes, sterile Trypsin/EDTA (Invitrogen, cat. no. 25300-054), warm

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

50-ml conical centrifuge tubes, sterile Assorted pipets, sterile 20-ml syringe, sterile Mixing cannula (Maersk Medical, cat. no. 500.11.012) Centrifuge Transfer pipet, sterile Hemacytometer Collagen I-coated 6-well plates (BD Biosciences Discovery Labware, cat. no. 356400) Inverted microscope Fine-tipped marker Prepared cloning cylinders (see Support Protocol 2) Forceps, sterile Pasteur pipets (Fisher Scientific, cat. no. 13-678-20C), sterilized 1.5- and 2.0-ml microcentrifuge tubes Collagen I coated 24-well tissue culture plate (see Support Protocol 1)

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25- and 75-cm2 vented tissue culture flasks (BD Biosciences Falcon), coated with rat tail collagen I (see Support Protocol 1) Additional reagents and equipment for counting viable cells using a hemacytometer and trypan blue exclusion (UNIT 1C.3) Dilute the sample 1. Aliquot 15 ml of UCB into each 50-ml conical centrifuge tube. UCB is collected with heparin or EDTA as an anticoagulant and transported to the laboratory at room temperature. For the most ideal colony formation, UCB should be processed within 2 hr of the infant’s delivery. If immediate processing is not feasible, UCB can be kept at room temperature with continuous rocking for up to 16 hr prior to isolation of MNCs. Colony numbers decline with increased time between delivery and processing.

2. Add 20 ml PBS to each tube of UCB and pipet several times to mix.

Isolate MNCs 3. Draw up 15 ml Ficoll into a 20-ml syringe and fit the syringe with a mixing cannula. 4. Place the tip of the mixing cannula at the bottom of the tube of diluted blood and carefully underlay 15 ml Ficoll. Remove all air from the syringe and mixing cannula prior to submerging the tip into the diluted blood, as air bubbles will disrupt the blood-Ficoll interface. If there are multiple tubes of diluted blood, Ficoll can be dispensed from 30- or 60-ml syringes.

5. Centrifuge the tubes 30 min at 740 × g, room temperature, with no brake. Handle all tubes carefully to maintain a clean blood-Ficoll interface. Bringing the centrifuge up to 740 × g slowly over the course of 1 to 2 min helps result in a cleaner buffy coat.

6. Using a transfer pipet, remove the hazy layer of MNCs that sits at the interface between the Ficoll and serum. Dispense the MNCs into a 50-ml conical tube containing 10 ml EBM-2 10:1. Following centrifugation, red blood cells will form a pellet, and MNCs will form a hazy buffy coat at the interphase between the clear Ficoll layer below and the yellow serum layer above (see Fig. 2C.1.1A). Care should be taken to collect all buffy coat MNCs while avoiding excess collection of the Ficoll layer or the serum layer. Immature red blood cells occasionally fail to separate cleanly from the MNCs (see Fig. 2C.1.1B). Make an effort to avoid collection of RBCs when removing the MNCs. The presence of RBCs, however, will not disrupt the culture of ECFCs.

7. Centrifuge the MNCs 10 min at 515 × g, room temperature with a high brake. 8. Carefully aspirate and discard the supernatant. Following this initial centrifugation, the pellet of cells is often loose. Care should be taken to avoid aspirating cells.

9. Gently tap the tube to loosen the pelleted cells and resuspend in 10 ml of EBM-2 10:1. If there are multiple tubes from the same sample, cell pellets can be serially combined at this point.

10. Repeat steps 7 to 9 one time. Somatic Stem Cells

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Figure 2C.1.1 Isolation of MNCs and culture of ECFCs. (A-B) Photographs of Ficoll separation of MNCs from adult peripheral blood (A) and UCB (B). Black arrows indicate the buffy coat of MNCs. White arrow indicates a layer of immature RBCs often present in UCB preparations. (C) Representative photomicrograph of ECFC colony 10 days following initial plating of UCB MNCs. Arrows indicate colony boundaries. Magnification 40×. (D) Representative photomicrograph of contaminating MSC colony arising from culture of UCB MNCs. Magnification 40×.

Assess viability 11. Remove 30 μl of the cell suspension and mix with 30 μl trypan blue. Count viable cells on a hemacytometer (UNIT 1C.3) and calculate the total number of MNCs in the sample. Total cell numbers at this point typically range between 0.8 and 3 × 106 MNCs per milliliter of whole blood processed. Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

12. Centrifuge the cell suspension 10 min at 515 × g, room temperature, with a high brake. Aspirate the supernatant.

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Seed MNCs for culture 13. Tap the tube to loosen the cell pellet and resuspend MNCs in cEGM-2 at 1.25 × 107 cells/ml. 14. Pipet 4 ml (5 × 107 MNCs) into each well of a 6-well tissue culture plate precoated with rat tail collagen I (see Support Protocol 1), and place in a 37◦ C, 5% CO2 humidified incubator. A seeding density in the range of 3 to 5 × 107 MNCs/well is ideal for ECFC colony formation from UCB.

Change medium 15. After 24 hr (day 1), slowly remove the medium from the well with a pipet. Medium is removed at a rate of 1 ml per 4 to 5 sec. Leave some liquid in the well to prevent drying of the plate surface.

16. Slowly add 2 ml cEGM-2 to the well. 17. Slowly remove the 2 ml of medium and add 4 ml cEGM-2 to the well. Return culture plates to the incubator. 18. After another 24 hr (day 2), refresh the medium by slowly removing the medium from the well with a pipet and adding 4 ml cEGM-2 to the well. Repeat medium change daily to day 7 and every other day thereafter. Each day from day 3 to day 7 of culture, medium is refreshed as in step 18. After day 7, medium can be changed every other day. After day 7, medium can be aspirated from the culture plates by vacuum, and medium can be added at a moderate rate of 1 ml per 2 to 3 sec. ECFC-derived colonies appear between day 4 and day 7 of culture as well circumscribed areas of cobblestone-appearing cells (see Fig. 2C.1.1C). Individual colonies can be isolated and expanded on day 7 to day 14. Cells can also be allowed to grow to 80% to 90% confluency before subculturing to start a polyclonal EC line. Rarely, mesenchymal stem cell (MSC) colonies can begin to grow from primary MNC cultures in these conditions (see Fig 2C.1.1D). MSC are identified as elongated, fibroblastlike cells and can quickly over-grow the ECFCs. ECFCs should be clonally isolated and subcultured immediately if MSC contamination is found.

Clone ECFCs 19. Visualize the ECFC colony using an inverted microscope and outline the colony boundaries with a fine-tipped marker on the underside of the well. 20. Prepare the necessary number of cloning cylinders (see Support Protocol 2 and Fig. 2C.1.2A). Use sterile forceps when handling cloning cylinders.

21. Aspirate the culture medium and wash the culture well two times, each time with 3 ml PBS. 22. After aspirating the final wash of PBS, place a cloning cylinder around each colony and press firmly against the plate using forceps. Leave a small amount of PBS in the culture well when aspirating the final wash to keep the surface from drying out. When placing the cloning cylinders around a colony, press firmly vertically to affix the cylinder to the plate. Avoid any side-to-side motions or sliding the cylinder on the culture surface as this disrupts the ECFC colony. Somatic Stem Cells

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Figure 2C.1.2 Isolation of ECFC colonies using cloning cylinders. (A) Photograph of a cloning cylinder prepared with vacuum grease. (B) Photograph of placement of a cloning cylinder for isolation of a single ECFC colony from a 6-well culture plate.

23. Using a Pasteur pipet fitted with a bulb, add 1 to 2 drops of warm trypsin/EDTA into each cloning cylinder (see Fig. 2C.1.2B). Perform steps 22 and 23 quickly to prevent cells from drying out.

24. Incubate plates for 1 to 5 min until the cells within the cylinder begin to ball up and detach.

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

25. When all the cells within the cylinder have balled up, place the tip of a Pasteur pipet containing 200 to 300 μl of cEGM-2 into the center of the cylinder and pipet up and down vigorously several times. Collect the entire volume into a microcentrifuge tube. Wash the area within the cylinder 1 to 3 more times with 200 to 300 μl cEGM-2 until all cells are collected into the microcentrifuge tube. 26. Seed the cells from each ECFC colony into one well of a 24-well tissue culture plate precoated with rat tail collagen I (see Support Protocol 1) in a total volume of 1.5 ml of cEGM-2 and culture in a 37◦ C, 5% CO2 humidified incubator for expansion.

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Propagate ECFCs ECFC-derived cells can be propagated as monoclonal cell lines, initiated via the cloning steps above. Alternatively, primary ECFC colonies can be allowed to grow to 80% to 90% confluency in the original culture plate prior to subculturing for the propagation of a polyclonal cell line. 27. When cells approach confluency, remove culture medium and wash the culture surface two times, each time with 500 μl PBS. For collection of a polyclonal cell line from the original 6-well plate, use 3 ml PBS for washes.

28. Aspirate the final wash of PBS and add trypsin/EDTA. Warm trypsin-EDTA in a 37◦ C water bath prior to use. Use 200 μl/well for 24-well plates, 400 μl/well for 6-well plates, 750 μl/25-cm2 flasks, and 1.5 ml/7-cm2 flasks.

29. Incubate at 37◦ C until the cells ball up and begin to detach. 30. Add cEGM-2 to the cells and collect into a 15- or 50-ml tube. Examine the culture surface under low (10 to 20×) magnification with an inverted microscope. If cells remain, wash the culture surface again with cEGM-2 and collect into the tube. Use 1 ml/well for 24-well plates, 2 ml/well for 6-well plates, 4 ml/25-cm2 flask, and 8 ml/75-cm2 flask. If cells remain on the culture surface following two washes with cEGM-2, repeat steps 27 to 30 to collect all remaining cells.

31. Obtain a viable cell count of an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). 32. Seed 3000 to 5000 cells/cm2 onto a collagen I coated tissue culture surface (see Support Protocol 1) in cEGM-2 medium. Use 500 μl cEGM-2/well for 24-well plates, 3 ml/well for 6-well plates, 5 ml/25-cm2 flasks, and 10 ml/75-cm2 flasks

33. Replace culture medium with fresh cEGM-2 every other day. Culture until cells approach 80% to 90% confluency before subculturing again. Cultures typically approach confluency in 3 to 7 days.

ISOLATION, CLONING, AND PROPAGATION OF ENDOTHELIAL COLONY-FORMING CELLS FROM HUMAN PERIPHERAL BLOOD

ALTERNATE PROTOCOL 1

Primary culture of ECFCs from adult peripheral blood can be achieved using the same protocol described for UCB-derived ECFCs (see Basic Protocol 1). An amount of 100 ml of venous blood is collected in heparin and MNCs are isolated and cultured exactly as described in Basic Protocol 1. ECFC colonies will appear between day 14 and day 28 of culture and can be clonally isolated and expanded. NOTE: The frequency of peripheral blood derived–ECFCs is ∼1 in 20 ml of blood (Ingram et al., 2004); therefore, it is necessary to begin with 80 to 100 ml of blood to ensure colony growth. NOTE: For peripheral blood, CPT vacutainers with heparin or citrate anticoagulant (BD Vacutainer) can be used as an alternative to Ficoll separation of MNCs. Collect blood in CPT vacutainers and centrifuge directly in the collection tubes according to the manufacturer’s instructions for initial separation of MNCs. After removing the MNCs from the CPT vacutainers, wash and plate cells exactly as described in Basic Protocol 1.

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BASIC PROTOCOL 2

PHENOTYPIC CHARACTERIZATION OF ECFCs: ENDOTHELIAL-SPECIFIC CELL SURFACE ANTIGEN EXPRESSION ECFCs arise from primary culture of MNCs. To confirm the endothelial phenotype of outgrowing cells and ensure the absence of any other contaminating cell type, ECFC cell lines should be examined by several methods. Pure cultures of ECFCs derived from UCB or peripheral blood uniformly express the EC-specific cell surface antigens, CD31, CD105, CD144, and CD146, but do not express the hematopoietic cell–specific antigen CD45 or the monocyte/macrophage marker, CD14. Functionally, ECFCs will ingest AcLDL (Alternate Protocol 2) and form a capillary tube-like network when placed in Matrigel (Alternate Protocol 3). Numerous other methods exist to confirm an endothelial phenotype including detection of surface and intracellular markers (Lin et al., 2000; Gulati et al., 2003; Bompais et al., 2004; Hur et al., 2004; Yoder et al., 2007) like von Willebrand Factor (vWF), KDR (VEGF-R2), Tie-2, and eNOS; functional binding of EC-specific lectins (Kalka et al., 2000; Dimmeler et al., 2001; Gulati et al., 2003; Hill et al., 2003; Yoder et al., 2007), UEA-1 and BS-1; and upregulation of VCAM-1 and ICAM-1 (Lin et al., 2000; Bompais et al., 2004). Investigators can consult the literature for method details. NOTE: Known endothelial cell lines, such as human umbilical vein endothelial cells (HUVEC) or human microvascular endothelial cells (HMVEC), can be used as a positive control for all phenotyping assays. These cells are commercially available from several vendors.

Materials ECFC cultures grown in 25- or 75-cm2 flasks (Basic Protocol 1) Trypsin-EDTA (Invitrogen, cat. no. 25300-054) Staining buffer (see recipe), ice cold Phosphate-buffered saline (PBS), without calcium and magnesium 0.4% (w/v) trypan blue solution Fc Block (Miltenyi, cat. no. 130-059-901) hCD31 antibody, FITC conjugated (BD Pharmingen, cat. no. 555445) hCD45 antibody, FITC conjugated (BD Pharmingen, cat. no. 555482) hCD14 antibody, FITC conjugated (BD Pharmingen, cat. no. 555397) hCD144 antibody, PE conjugated (eBioscience, cat. no. 12-1449-80) hCD146 antibody, PE conjugated (BD Pharmingen, cat. no. 550315) hCD105 antibody, PE conjugated (Invitrogen, cat. no. MHCD10504) Ms IgG1,κ antibody, FITC conjugated (BD Pharmingen, cat. no. 555748) Ms IgG1,κ antibody, PE (BD Pharmingen, cat. no. 559320) Ms IgG2a,κ FITC conjugated (BD Pharmingen, cat. no. 555573) Fixing buffer (see recipe), ice cold 15- or 50-ml conical centrifuge tubes, sterile Hemacytometer 12 × 75–mm polystyrene round bottom tubes (BD Falcon, cat. no. 352008) FACSCalibur flow cytometer (Becton Dickinson) or equivalent Flow cytometry analysis software (e.g., Cellquest; Becton Dickinson or FlowJo; Tree Star)

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

Additional reagents and equipment for detaching cells with tryspin/EDTA (Basic Protocol 1) and counting viable cells using a hemacytometer and trypan blue exclusion (UNIT 1C.3)

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Current Protocols in Stem Cell Biology

Collect cells for staining 1. Detach cells with trypsin/EDTA according to the steps described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 29). Use early passage (1 to 3) ECFC cultures, which are 50% to 80% confluent. One 75-cm2 flask of ECFCs should yield sufficient cell numbers for subsequent staining for cell surface antigen expression.

2. Add 4 ml staining buffer to the cells and collect into a 15-ml tube. If cells remain on the culture surface, wash 1 to 2 more times with 4 ml staining buffer and collect the washes into the 15-ml tube. 3. Obtain a viable cell count from an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). 4. Prepare ten 12 × 75–mm polystyrene round bottom tubes and aliquot up to 106 cells/tube. Bring total volume of each tube up to 3 ml with staining buffer. FACS tubes should be clearly labeled with the antibody used. Prepare one tube for each primary antibody and isotype listed above. Also prepare one tube for unstained cells to serve as a negative control. 106 cells are not required for each test. The authors typically prepare 0.5 to 1 × 105 cells/tube, for the eventual collection of 2 × 104 analyzed events on a flow cytometer. Up to 106 cells can be stained in each tube using the same amount of antibody.

5. Centrifuge tubes 10 min at 515 × g, 4◦ C. Pour off supernatant and blot residual supernatant onto a paper towel. When decanting the supernatant, invert the tube only once and take care not to disrupt the pellet of cells. While the tube is inverted, blot the rim of the tube once on a paper towel to remove excess liquid.

Stain with primary antibody 6. Resuspend cells in 100 μl staining buffer. 7. Add 10 μl Fc block to each tube. Mix gently and place on ice for 10 min. 8. Add the appropriate amount of one primary antibody or isotype to each tube. Swirl tubes gently and leave on ice 30 min protected from light. Follow the manufacturer’s recommendation for the amount of antibody to be used for each test. Some antibodies may require titrating to determine the optimal concentration. One FITC-conjugated antibody and one PE-conjugated antibody can be used to costain the cells in the same tube, provided that appropriate single-color controls are prepared as compensation controls for the flow cytometer. For more information on immunophenotyping via flow cytometry see Baumgarth and Roederer (2000), Shapiro (2003), and Perfetto et al. (2006).

Wash the cells 9. Add 3 ml cold staining buffer to each tube and centrifuge 10 min at 515 × g, 4◦ C. 10. Decant supernatant and blot residual supernatant onto a paper towel. Tap the tube to loosen each pellet. 11. Repeat steps 9 and 10 one time.

Fix the cells 12. Resuspend each pellet in 300 μl fixing buffer. Store fixed samples at 4◦ C, protected from light for at least 24 hr prior to analysis, on a flow cytometer. If cells will be analyzed on a flow cytometer immediately, cells can be suspended in staining buffer rather than fixing buffer at this step. Fixed cells to be analyzed at a later time should be stored at 4 ◦ C protected from light.

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Figure 2C.1.3 Representative phenotypic analysis of UCB-derived ECFCs. (A) Assessment of cell surface expression of endothelial cell specific antigens CD31, CD105, CD144, and CD146; hematopoietic cell specific CD45; and monocyte/macrophage specific CD14 on passage 3 UCBderived ECFCs. Filled black histograms represent antigen staining. Negative isotype controls are overlaid in red. (B) Passage 2 UCB-derived ECFCs following 4 hr incubation with DiI-AcLDL (red). Nuclei are counterstained with DAPI (blue). Magnification 100×. (C) Passage 3 UCB-derived ECFCs 24 hr post seeding onto Matrigel coated wells. Magnification 40×.

Analyze the cells 13. Analyze samples on a flow cytometer within 7 days to determine the percentage of cells that stain positively for each antigen. Fixed samples are analyzed directly in fixing buffer. Collection of 2 × 104 events is sufficient to discern surface antigen expression. ECFCs will be uniformly positive for CD31, CD105, CD144, and CD146, but negative for CD45 and CD14 compared to the corresponding isotype controls (see Fig. 2C.1.3A). ALTERNATE PROTOCOL 2

PHENOTYPIC CHARACTERIZATION OF ECFCs: UPTAKE OF AcLDL

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

Additional Materials (also see Basic Protocol 2) Collagen I coated 24-well tissue culture plate (see Support Protocol 1) DiI-complexed acetylated low density lipoprotein from plasma (DiI AcLDL; Invitrogen, cat. no. L-3484)

A functional assay for ECFC is the uptake of acetylated LDL. ECFCs, which ingest the DiI-labeled AcLDL described in this protocol, can be detected by their red fluorescence.

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EBM-2 10:1 (see recipe) Inverted fluorescence microscope with a rhodamine filter Additional reagents and equipment for preparing a culture of ECFCs (Basic Protocol 1) 1. Prepare a culture of ECFCs in a collagen I-coated 24-well tissue culture plate as described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 32). Allow cells to incubate and adhere 8 to 24 hr prior to assaying for AcLDL uptake. 2. Remove culture medium and wash wells two times, each time with 500 μl PBS. 3. Add 500 μl EBM-2 10:1 containing 10 μg/ml DiI AcLDL. Incubate 4 to 6 hr. 4. Remove medium containing DiI AcLDL and wash wells two times, each time with 500 μl PBS. Add 500 μl EBM-2 10:1 to each well. 5. Examine cells for uptake of DiI AcLDL using an inverted fluorescence microscope. Cells which have ingested AcLDL will fluoresce red. ECFCs will ingest AcLDL and fluoresce red (see Fig. 2C.1.3B). Cell nuclei can be counterstained with a nuclear dye, for example DAPI, to identify all cells under fluorescent microscopy. As an alternative to visualizing AcLDL uptake with a fluorescence microscope, cells can be detached, resuspended in FACS staining buffer, and run on a flow cytometer. Other fluorescent conjugates (e.g., FITC or AlexaFluor 488) can be used instead of DiI to visualize AcLDL uptake.

PHENOTYPIC CHARACTERIZATION OF ECFCs: MATRIGEL LATTICE FORMATION

ALTERNATE PROTOCOL 3

Another functional characteristic of ECFCs is their ability to form networks of capillarylike structures when cultured on Matrigel.

Additional Materials (also see Basic Protocol 2) cEGM-2 (see recipe) Additional reagents and equipment for preparing a Matrigel-coated tissue culture plate (Support Protocol 5) and obtaining a viable cell count using a hemacytometer and trypan blue exclusion (UNIT 1C.3) 1. Prepare a Matrigel-coated tissue culture plate (see Support Protocol 5). 2. Detach and collect cells according to the steps described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 30). 3. Centrifuge cells 10 min at 515 × g, room temperature, and discard the supernatant. 4. Tap the tube to loosen the cell pellet and resuspend in 5 to 10 ml cEGM-2. Volume is dependent on the number of cells collected. Cell concentration should be at least 2.5 × 104 cells/ml.

5. Obtain a viable cell count of an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). 6. Calculate the volume of cell suspension necessary for 5000, 7500, and 104 total cells. If the volume of cell suspension needed is >200 μl, centrifuge cells again and resuspend in a smaller volume of cEGM-2. Perform another cell count and calculate volumes again.

7. Mix the cell suspension thoroughly and seed 5000 cells to each of three Matrigelcoated wells. Add cEGM-2 to each well to make up the total volume of medium to 200 μl.

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8. Repeat step 7 for seeding 7500 and 104 cells/well. 9. Incubate plates and examine using an inverted microscope under 20 to 100× magnification every 2 hr for capillary-like tube formation. ECFCs will begin to migrate and form a lattice-like network within a few hours and will continue to elongate to form a continuous network at optimal seeding densities (see Fig. 2C.1.3C). Viable cells, which fail to form tubes at or before 24 hr, are not ECs. Capillarylike networks can be quantitated, if desired, using software, such as ImageJ (available at http://rsb.info.nih.gov/ij/) to measure vessel length. BASIC PROTOCOL 3

TRANSPLANTATION OF ECFCs INTO MICE We have determined that the most stringent means to verify the functionality of ECFCs is to assess their ability to contribute to de novo vasculogenesis. This protocol describes how to cast ECFCs in a collagen-fibronectin matrix, implant and harvest the cellularized grafts, and assess vasculogenesis by quantifying the density of blood vessels within the implant. 2 × 106 ECFCs are cast into a fibronectin-collagen matrix and allowed to form a primitive capillary network overnight. Cellularized gels are bisected and implanted into the flank of a NOD/SCID mouse (106 ECFCs/implant). One graft can be implanted on each side of a mouse’s abdomen allowing for placement of an internal control and test graft in each animal. NOTE: This protocol is written for the casting of one 1-ml gel, which will be bisected to yield 2 implants. To perform this on a larger scale, on multiple mice, multiply all volumes by the appropriate factor.

Materials Fetal bovine serum (FBS; Hyclone, cat. no. SH30070.03) EBM-2 10:1 (see recipe), ice cold 7.5% (w/v) sodium bicarbonate (Sigma, cat. no. S8761), sterile and ice cold 1 N NaOH, sterile and ice cold 1 M HEPES (Lonza, cat. no. 17-737E), ice cold 1 mg/ml fibronectin (Millipore, cat. no. FC10-10MG), ice cold Rat tail collagen type I, (BD Biosciences Discovery Labware, cat. no. 354236), ice cold cEGM-2 (see recipe), warm ECFC cultures grown in 25- or 75-cm2 flasks (Basic Protocol 1) Trypsin/EDTA (Invitrogen, cat. no. 25300-054) Phosphate-buffered saline (PBS), without calcium and magnesium 0.4% (w/v) trypan blue solution (Sigma, cat. no. T8154) Immunodeficient mice, 8- to 12-weeks-old (see Critical Parameters) Isoflurane inhalant Alcohol pads or 70% ethanol Zinc fixative (BD Biosciences, cat no. 550523)

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

37◦ C water bath Hemacytometer 15- or 50-ml conical centrifuge tubes, sterile Micropipettor with a 1-ml tip 12-well tissue culture plate Thin surgical spatula (e.g., FST, cat. no. 10091-12), sterile Fine iris scissors, sterile

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Electric shears Smooth forceps (2), sterile Sharp iris scissors, sterile Blunt-end iris scissors Light microscope with an eyepiece micrometer 5-0 polypropylene suture on a cutting needle Glass slides Additional reagents and equipment for obtaining a viable cell count using a hemacytometer and trypan blue exclusion (UNIT 1C.3), isoflurane anesthesia (UNIT 1B.4), euthanizing the mouse (Donovan and Brown, 2006), paraffin-embedding the gel (Bancroft and Gamble, 2002), and staining with hematoxylin and eosin (Bancroft and Gamble, 2002) or anti–human CD31 or anti–mouse CD31 (Support Protocol 7) to visualize the vasculature within the gel Prepare reagents 1. Calculate the total volume (ml) of gel material needed to cast the desired number of gel implants using: Vtot = 1.2 ml × (no. of gels) + 2 ml 1.2 ml is the volume of gel material and cells that are prepared to make each 1-ml gel. Each 1-ml gel is later bisected to yield two implants.

Calculate the volume of each component needed to prepare the gel material using the formulas in Table 2C.1.1. 2. Aliquot a small working stock (∼1.2× the volume of each component calculated in step 1) of each of the following gel components and place on ice: FBS, EBM-2 10:1, sodium bicarbonate, NaOH, HEPES, fibronectin, and collagen I. Aliquot a working stock of cEGM-2 and warm in a 37◦ C water bath.

Prepare cells 3. Detach cells with trypsin/EDTA according to the steps described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 30). If cells detach from the culture surface but remain adherent to each other, disrupt mechanically by pipetting up and down several times in the presence of trypsin/EDTA to ensure a single-cell suspension.

4. Obtain a viable cell count of an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). Table 2C.1.1 Formulas for the Calculation of Reagent Volumes Used in Casting Cellularized Gel Implants

Reagent

Stock conc.

Final conc.

HEPES

1M

25 mM

VHEPES = 25 μl/ml × Vtot

Sodium bicarbonate

7.5%

1.5 mg/ml

VNaBicarb = 20 μl/ml × Vtot

FBS

100%

10%

VFBS = 100 μl/ml × Vtot

Fibronectin

1 mg/ml

100 μg/ml

VFN = 100 μl/ml × Vtot

Collagen I

Variable

1.5 mg/ml

VColl = 1.5 mg/ml × Vtot / (Collagen stock conc. in mg/ml)

EBM-2 10:1

Calculation

VEBM2 = 0.7 × Vtot – (VHEPES + To bring solution volume to 0.7 × Vtot VNaBicarb + VFBS + VFN + VColl )

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5. For each gel, aliquot 2.4 × 106 cells into a 50-ml conical tube. 2.4 × 106 cells are used to cast one 1-ml gel containing 2 × 106 ECFCs/ml. This gel is later bisected to yield two implants containing 106 ECFCs. If multiple implants with the same cells are needed, cells can be combined in the same 50-ml conical tube at this step.

6. To pellet cells centrifuge 10 min at 515 × g, room temperature.

Prepare cellularized gel implants 7. While the cells are centrifuging, prepare the gel matrix solution by adding calculated volume of each component to an ice-cold 50-ml conical tube in this order: HEPES, sodium bicarbonate, EBM-2 10:1, FBS, fibronectin, and collagen I. Mix thoroughly. 8. Add 1 N NaOH in μl amounts, while monitoring the pH until the solution reaches pH 7.4. Keep solution on ice. All reagents and tubes must be ice cold. Approximately 3 μl of 1 N NaOH is used per 1 ml of gel solution to approach pH 7.4. The correct pH is critical for proper polymerization of the gel.

9. After cells have been centrifuged, discard the supernatant and resuspend the ECFC pellet to 360 μl in warm cEGM-2. The cell pellet typically consumes a volume of 50 to 100 μl. Cells are resuspended to a total volume of 360 μl, including the cell volume. It is critical at this point that the cells are dispersed into a single-cell suspension with no aggregates.

10. Using a micropipettor with a 1-ml tip, add 840 μl gel solution to each conical tube of suspended cells. Slowly mix until cells are thoroughly suspended in the gel solution. Adjust the micropipettor to 1 ml and aliquot 1 ml of cellularized gel solution to one well of a 12-well tissue culture plate. 11. Incubate plate for 20 to 30 min until the gel polymerizes. 12. Gently cover the gel with 2 ml warm cEGM-2 and incubate overnight Following 16 to 24 hr of incubation, ECFCs will form a dense capillary-like network within the gel matrix.

Implant gels 13. In the surgical facility, immediately prior to implantation, bisect the gel implant by carefully lifting it from the culture dish with a thin surgical spatula and cutting it in half with fine iris scissors. Return gel pieces to the culture well containing medium until implantation. 14. Administer isoflurane anesthesia. Refer to UNIT 1B.4, Support Protocol 2, for mouse anesthesia. See Figure 1B.4.1A for the setup of the anesthesia unit.

15. Using electric shears, shave the lower part of the abdomen and clear loose hair from the surgical site. Thoroughly clean the surgical site with alcohol pads or 70% ethanol. 16. Using forceps, pinch a skin fold in the lower quadrant of the abdomen and make an ∼5-mm incision into the skin fold with sharp iris scissors (see Fig. 2C.1.4A), exposing the subcutaneous space between the skin and abdominal muscle. When making the incision, take care not to cut or tear the soft abdominal muscle. Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

17. Carefully dissect the dermal layer from the abdominal muscle by inserting closed blunt-end iris scissors under the skin and gently opening the scissors to create a pocket (∼15 × 20–mm) leading superior into the upper abdominal quadrant (see Fig. 2C.1.4B).

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Figure 2C.1.4 Surgical implantation and harvest of cellularized gel grafts. (A-B) Illustration of initial incision and creation of subdermal pocket prior to implantation of cellularized gel. (C) Illustration of bilateral cellularized gel placement prior to closure of incisions. Black arrows indicate gel location. (D) Illustration of cellularized gel appearance in situ at the time of harvest. White arrows indicate gel location. (E) Representative photograph of a cellularized gel at the time of harvest. This gel contained UCB-derived ECFC and ADSCs and was harvested 14 days after implantation. Vascularization within the gel is visible by the red coloration.

18. With one set of forceps, pinch and lift the dermal layer just caudal to the incision to open the pocket. Using a second set of smooth forceps, lift one piece of the bisected gel from the culture dish and insert into the dissected pocket. Visualize for proper placement (see Fig. 2C.1.4C). Most gels do not retain their original semi-circular shape during implantation.

19. Repeat steps 16 to 18 to implant the second gel on the other side of the mouse’s abdomen. 20. Close each incision with 2 or 3 stitches using 5-0 polypropylene suture on a cutting needle. Visualize gels to ensure that they remain deep inside the pocket during closure of the incision. If gels will be implanted into multiple animals, instruments and unused suture lengths can be held in sterile PBS between surgeries for each animal.

21. Label cage cards with details of the procedure. 22. Administer post-surgical monitoring and analgesia according to institutional requirements and protocols.

Harvest gel implants Cellularized gel implants can be harvested and examined for the formation of vasculature as early as 48 hr after implantation. To ensure the ability of the vasculature to mature, gels should reside in the animal for 7 days. 23. Euthanize the mouse (Donovan and Brown, 2006) in accordance with local regulations. Euthanasia can be achieved by CO2 asphyxiation followed by cervical dislocation.

24. Swab the abdominal area with 70% ethanol or alcohol pads. Somatic Stem Cells

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25. Using scissors, cut the abdominal skin caudal to the original incision line. Carefully dissect the skin by excising a flap of skin caudal to the probable location of the gel. Take care to dissect the skin away from the gel, leaving it adhered to the abdominal muscle (see Fig. 2C.1.4D,E). Dissect carefully to ensure that the gel remains intact. Often the gel may migrate slightly from the original implantation site by a few millimeters. The gel may appear white, faint pink, or deep red depending on the extent of vascularization.

26. Excise the implant by cutting circumferentially around the gel and place in zinc fixative. When excising the gel, include a boundary of mouse tissue to serve as an internal control of vasculature. Other standard fixatives (e.g., formalin) may be appropriate depending on the antibody that will be used to stain the specimens. Experimenters should refer to the antibody manufacturer’s recommendations for fixation methods.

27. Allow gel tissues to fix 1 to 2 hr at room temperature.

Embed and stain immunohistochemically 28. Paraffin-embed the gel according to standard histochemical protocols (see Bancroft and Gamble, 2002). 29. Prepare 5-μm sections on glass slides. To achieve accurate representation of overall vascular density within the specimen, multiple sections ∼100 μm apart should be prepared.

30. Stain sections with hematoxylin and eosin (see Bancroft and Gamble, 2002), anti– human CD31, or anti–mouse CD31 (see Support Protocol 7) to visualize the vasculature within the gel. Staining with anti–mouse and anti–human CD31 is necessary to confirm the origin of vasculature within the gel.

31. Visualize stained tissue sections under a light microscope (Fig. 2C.1.5A,B).

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

Figure 2C.1.5 Immunohistochemical staining of cellularized gel implants for quantitation of vascularization. (A) Representative photomicrograph of cellularized (ECFCs only) gel implant and surrounding mouse tissue stained with H&E (blue and pink) and anti–human CD31 (brown). Black arrows indicate RBC perfused, anti–human CD31+ vessels within the gel implant. Magnification 20×. (B) Representative photomicrograph of cellularized (ECFCs and ADSCs) gel implant and surrounding mouse tissue stained with H&E (blue and pink). Black arrows indicated RBC-perfused vessels within the gel implant. Magnification 100×.

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Quantify vascularization The extent of vascularization within the gel implant is best expressed as average number of red blood cell-containing anti–human CD31-positive vessels per mm2 of gel implant. 32. Count the number of anti–human CD31-positive vessels, which contain red blood cells under 40× magnification of a light microscope (Fig. 2C.1.5A,B). 33. Measure the dimensions of the gel implant and calculate the area (in mm2 ) of the implant sample. 34. Calculate the number of red blood cell–containing vessels/mm2 by dividing the count from step 32 by the area calculated in step 33. For an accurate representation of vasculature within the gel implant, at least four separate planes of the implant should be scored and averaged.

TRANSPLANT OF MIXED CELL IMPLANTS INTO MICE Cellularized grafts containing ECFCs only tend to yield vessels which are unstable and prone to microaneurysm. Adipose-derived stem cells (ADSC) can be cocultured at a 1:4 (ADSC:ECFC) ratio in the gel implants to establish more stable vasculature.

ALTERNATE PROTOCOL 4

Additional Materials (also see Basic Protocol 3) Adipose-derived stem cells (ADSC; Lonza, cat. no. PT-5006), grown in 25- or 75-cm2 flasks (see Support Protocol 6) Prepare mixed cell gel implants 1. Follow steps 1 to 4 of Basic Protocol 3 to collect and count ECFCs. 2. Collect ADSCs in cEGM-2 and obtain a viable cell count according to steps 27 to 31 of Basic Protocol 1. 3. For each gel, aliquot 1.92 × 106 ECFCs and 4.8 × 105 ADSCs to a 50-ml conical tube. 2.4 × 106 total cells are used to cast one 1-ml gel containing 1.6 × 106 ECFCs and 4 × 105 ADSCs/ml. This gel is later bisected to yield 2 implants containing 106 total cells. If multiple implants with the composition are needed, cells can be combined in the same 50-ml conical tube at this step.

4. Follow steps 6 to 12 exactly as in Basic Protocol 3. 5. For implanting and harvesting gel implants and quantifying vasculature, follow steps exactly as outlined in Basic Protocol 3, steps 13 to 34.

PREPARATION OF COLLAGEN-COATED TISSUE CULTURE SURFACES ECFCs are cultured and propagated on culture surfaces coated with type I collagen.

SUPPORT PROTOCOL 1

Materials Collagen I solution (see recipe) Phosphate-buffered saline (PBS), without calcium and magnesium Tissue culture–treated plates or flasks Pipets, sterile Pasteur pipets, sterile 37◦ C incubator 1. Place 1 ml collagen I solution in each well of a 6-well tissue culture–treated plate. Use 300 μl/well for 24-well plates, 4 ml/25-cm2 flasks, and 9 ml/75-cm2 flasks.

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2. Incubate 90 min to overnight at 37◦ C. 3. Remove the collagen I solution and wash surface two times, each time with PBS. Use 500 μl/well for 24-well plates, 5 ml/25-cm2 flask, and 10 ml/75-cm2 flask.

4. Use immediately for cell cultures. SUPPORT PROTOCOL 2

PREPARATION OF CLONING CYLINDERS Efficient cloning of primary ECFC colonies requires the use of sterile cloning cylinders to create a barrier from the surrounding MNC culture. Prepare cloning cylinders with vacuum grease just prior to use.

Materials Vacuum grease (Dow Corning, cat. no. 1658832) Glass dish Forceps, sterile Cloning cylinders, sterile (Fisher Scientific, cat. no. 07-907-10) 10-cm petri dish, sterile 1. Spread a dime-sized amount of vacuum grease into a thin layer in glass dish. 2. Autoclave, sterilize, and cool completely. 3. Using forceps, remove a cloning cylinder from its packaging and dip the bottom surface into the vacuum grease to coat. Apply the minimum amount of grease necessary to coat the bottom surface and form a good seal with a culture plate. Excess grease will interfere with the collection of cells (see Fig. 2C.1.2A).

4. Lightly set the prepared cylinder, greased-side down, in a petri dish until use. Prepare cloning cylinders just prior to use. SUPPORT PROTOCOL 3

CRYOPRESERVATION OF ECFCs ECFCs can be expanded in culture for a limited number of passages, so it is necessary to cryopreserve cell lines as a stock for future experiments. Cryopreserve ECFCs derived from UCB and peripheral blood using the same protocol.

Materials ECFC cultures grown in 25- or 75-cm2 flasks (Basic Protocol 1) Trypsin/EDTA (Invitrogen, cat. no. 25300-054) EBM-2 10:1 (see recipe) 0.4% (w/v) trypan blue solution (Sigma, cat. no. T8154) Freezing medium (see recipe), ice cold Phosphate-buffered saline (PBS), without calcium and magnesium

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15- or 50-ml conical centrifuge tubes, sterile Hemacytometer Cryovials Cryogenic-controlled rate freezing container (Nalgene) or insulated polystyrene foam box Liquid nitrogen storage container Additional reagents and equipment for performing a viable cell count using a hemacytometer and trypan blue exclusion (UNIT 1C.3)

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1. Detach cells with trypsin/EDTA according to the steps described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 29). Cells should be 60% to 80% confluent at the time of collection for cryopreservation.

2. Add EBM-2 10:1 to the cells and collect into a 15- or 50-ml tube. Use 4 ml/25-cm2 flask or 8 ml/75-cm2 flask. Examine the culture surface under low (10 to 20×) magnification with an inverted microscope. If cells remain, wash the culture surface again with EBM-2 10:1 and collect into the tube.

3. Obtain a viable cell count of an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). 4. To pellet cells, centrifuge 10 min at 515 × g, 4◦ C. Discard supernatant and tap the tube to loosen the cell pellet. 5. Gently resuspend the cell pellet in cold freezing medium at 0.5 to 1 × 106 cells/ml. 6. Aliquot 1 ml of the cell suspension into each cryovial. Cryovials should be clearly labeled with the cell line name, date, number of cells, and passage number.

7. Transfer cryovials into the freezing container and place at −80◦ C overnight. 8. The next day, transfer cryovials to a liquid nitrogen storage container.

THAWING CRYOPRESERVED ECFCs Cryopreserved ECFCs can be stored long term and thawed for continued propagation and use in various assays.

SUPPORT PROTOCOL 4

Materials Cryopreserved ECFCs in cryovials (Support Protocol 3) cEGM-2 (see recipe) 0.4% (w/v) trypan blue solution (Sigma, cat. no. T8154) 37◦ C water bath 15- or 50-ml conical centrifuge tubes, sterile Hemacytometer 25- and 75-cm2 vented tissue culture flasks (BD Falcon), coated with rat tail collagen I (see Support Protocol 1) Additional reagents and equipment for obtaining a viable cell count using a hemacytometer and trypan blue exclusion (UNIT 1C.3) 1. Remove cryovials from liquid nitrogen storage and place immediately into a 37◦ C water bath until slushy. 2. Pour cells into a 15-ml tube containing 9 ml of warm cEGM-2 medium. 3. Mix gently and obtain a viable cell count of an aliquot using a hemacytometer and trypan blue exclusion (UNIT 1C.3). 4. Seed 3000 to 5000 cells/cm2 onto a collagen I–coated tissue culture surface in cEGM-2 medium. Use 5 ml cEGM-2/25-cm2 flasks and 10 ml/7-cm2 flasks.

5. Incubate and allow cells to adhere for 4 hr, then remove medium and replace with fresh cEGM-2. Continue to culture cells according to the corresponding steps described for propagation of ECFCs (see Basic Protocol 1, steps 27 to 33).

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SUPPORT PROTOCOL 5

COATING 96-WELL PLATES WITH MATRIGEL Matrigel coating on 96-well plates is prepared just prior to plating a capillary-tube forming assay (see Alternate Protocol 3).

Materials Matrigel (BD Biosciences, cat. no. 356234) Pipet tips Pipettor 96-well, flat-bottomed tissue culture plate 1. Completely thaw Matrigel in the refrigerator overnight. 2. On ice, pipet 30 μl Matrigel into the necessary number of wells of the 96-well plate. Matrigel must be kept on ice at all times to prevent polymerization. If Matrigel begins to solidify and build up in the pipet tip, use a new tip or use cold tips. Avoid air bubbles when adding Matrigel to the wells as they will obscure visualization of tube formation.

3. Incubate culture plate for 10 min at 37◦ C to allow Matrigel to polymerize. Use immediately. Plates must be used immediately following preparation, otherwise the Matrigel coating will begin to dry out. If coated wells are not to be used immediately, add 30 to 50 μl medium to the wells to prevent drying out. SUPPORT PROTOCOL 6

ADSC CULTURE ADSCs are used as a carrier cell to support stable vessel formation within cellularized collagen/fibronectin gel implants. Culture ADSCs exactly as instructed by the manufacturer.

Materials Adipose-derived stem cells (ADSC; Lonza, cat. no. PT-5006) ADSC-GM (see recipe; Lonza) Trypsin/EDTA 0.4% (w/v) trypan blue solution (Sigma, cat. no. T8154) 25- or 75-cm2 tissue culture–treated flasks Hemacytometer Assorted pipets 15- or 50-ml conical centrifuge tubes, sterile Additional reagents and equipment for obtaining a viable cell count using a hemacytometer and trypan blue exclusion (UNIT 1C.3) 1. Thaw ADSCs according to the manufacturer’s instructions and seed at 5000 cells/cm2 onto a tissue culture–treated surface in ADSC-GM. Use 5 ml/25-cm2 flask and 10 ml/75-cm2 flask.

2. Refresh medium every 3 to 4 days. 3. When cells near 90% confluency, detach with trypsin/EDTA as described in Basic Protocol 1 and collect in ADSC-GM. Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

4. Obtain a viable cell count of an aliquot using trypan blue exclusion (UNIT 1C.3). 5. Seed cells into new culture flasks at 5000 cells/cm2 .

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CD31 IMMUNOHISTOCHEMICAL STAINING Anti–human or anti–mouse CD31 staining is performed to identify and confirm the origin of vasculature within cellularized collagen-fibronectin grafts following explantation.

SUPPORT PROTOCOL 7

Materials Zinc-fixed, paraffin-embedded 5-μm tissue sections on glass slides Xylenes Ethyl alcohol Phosphate-buffered saline (PBS) with calcium and magnesium Retrieval solution (Dako, cat. no. S236984) Blocking solution/diluent (Vector Labs, cat. no. SP-5050) Anti–mouse CD31 (clone mec13.3, available from various suppliers) Anti–human CD31 (clone JC70/A, Dako) Universal LSAB2 link-biotin kit (Dako, cat. no. K0675) DAB solution (Dako, cat. no. K3467) Coplin jars Additional reagents and equipment for deparaffinizing and hydrating tissue sections through a series of xylenes and serial alcohol dilutions (Bancroft and Gamble, 2002) 1. In Coplin jars, deparaffinize and hydrate tissue sections through a series of xylenes and serial alcohol dilutions using standard histology protocols (see Bancroft and Gamble, 2002). 2. Immerse slides in retrieval solution for 20 min at 95◦ C to 99◦ C. Allow slides to cool to room temperature. Rinse slides 1 to 2 times, each time in Coplin jars containing PBS. 3. Immerse slides in blocking solution for 15 min at room temperature. No rinsing is necessary following this antigen blocking step.

4. Incubate the slides with CD31 diluted in blocking solution for 30 min at room temperature. Typical primary antibody concentrations range from 1:100 to 1:4000 and should be determined by the researcher to ensure optimal staining.

5. Incubate slides with the secondary antibody and streptavidin-HRP according to the LSAB2 link-biotin kit. 6. Develop with DAB solution.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

ADSC-GM ADSC Basal Medium supplemented with the entire ADSC Bullet kit (Lonza, cat. no. PT-4505), 10% (v/v) FBS, and 1% (v/v) penicillin (10,000 U/ml)/streptomycin (10,000 μg/ml)/amphotericin (25 μg/ml). Store up to 1 month at 4◦ C.

Collagen I solution Dilute 0.575 ml of glacial acetic acid (17.4 N; Fisher, cat. no. A38-500) in 495 ml of sterile distilled water (0.02 N final concentration). Sterile filter the dilute acetic acid continued

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with a 0.22-μm vacuum filtration system (Millipore, cat. no. SCGPU05RE). Add 25 mg rat tail collagen I (BD Biosciences Discovery Labware, cat. no. 354236) to the dilute acetic acid to a final concentration of 50 μg/ml. The volume of collagen added will vary depending on the collagen stock concentration. Store up to 1 month at 4◦ C.

Complete EGM-2 (cEGM-2) EGM-2 (Lonza, cat. no. CC-3162) supplemented with the entire growth factor bullet kit, 10% (v/v) fetal bovine serum (FBS; Hyclone), and 1% (v/v) penicillin (10,000 U/ml)/streptomycin (10,000 μg/ml)/amphotericin (25 μg/ml; Invitrogen, cat. no. 15240-062). Store up to 1 month at 4◦ C.

EBM-2 10:1 EBM-2 (Lonza, cat. no. CC-3156) supplemented with 10% (v/v) fetal bovine serum (FBS; Hyclone, cat. no. SH30070.03) and 1% (v/v) penicillin (10,000 U/ml)/streptomycin (10,000 μg/ml)/amphotericin (25 μg/ml, Invitrogen; cat. no. 15240-062). Store up to 1 month at 4◦ C.

Fixing buffer Phosphate-buffered saline (PBS) with 1% (v/v) formaldehyde (Tousimis, cat. no. 1008B) Store up to 2 weeks at 4◦ C

Freezing medium 95% (v/v) fetal bovine serum (FBS; Hyclone) 5% (v/v) DMSO, sterile filtered Prepare fresh Staining buffer Phosphate-buffered saline (PBS) supplemented with 2% (v/v) fetal bovine serum (FBS) Store at 4◦ C for 2 weeks

COMMENTARY Background Information

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

Circulating EPCs are widely studied as biomarkers to assess risk and severity of cardiovascular disease and as cell-based therapy for several human cardiovascular disorders. Three major methods exist for culture of circulating EPCs from blood MNCs (Prater et al., 2007). One method, originally introduced by Asahara et al. (1997), has been subsequently modified (Ito et al., 1999; Hill et al., 2003) and can now be performed using a commercially available kit (Endocult, StemCell Technologies). In this method, MNC cultures yield discrete, adherent colonies, termed colony-forming unit-ECs (CFU-ECs), by day 5 to 9. CFU-ECs display some phenotypic and functional characteristics of endothelial cells, including expression of cell surface antigens, CD31, CD105, CD144, CD146, vWF, and KDR (VEGF-

R2) and uptake of AcLDL. However, they also express hematopoietic-specific antigens CD45 and CD14 and display nonspecific esterase and phagocytic capabilities consistent with monocyte/macrophages (Yoder et al., 2007) and cannot be propagated long term in culture. A second method employs a similar approach to identify adherent circulating angiogenic cells (CACs) from MNCs following 4 days of culture in endothelial specific conditions (Kalka et al., 2000; Dimmeler et al., 2001). Likewise, CACs resemble ECs phenotypically (Asahara et al., 1999; Kalka et al., 2000; Dimmeler et al., 2001), but have also proven to be enriched for hematopoieticderived monocyte/macrophages (Hassan et al., 1986; Rehman et al., 2003; Schmeisser et al., 2003; Ziegelhoeffer et al., 2004). Although less studied, we and others have identified ECFCs (Ingram et al., 2004), which

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are also referred to as blood outgrowth endothelial cells (BOECs; Lin et al., 2000; Gulati et al., 2003; Bompais et al., 2004; Hur et al., 2004), via a third method of culture of MNCs from human peripheral blood. ECFCs express cell surface antigens, CD31, CD105, CD144, CD146, vWF, and KDR, uptake of AcLDL, upregulate VCAM-1, and form capillary-like tubes when plated on Matrigel (Lin et al., 2000; Gulati et al., 2003; Hur et al., 2004; Ingram et al., 2004; Yoder et al., 2007). Additionally, ECFCs are organized in a hierarchy of progenitor stages that vary in proliferative potential and can be identified in clonal plating conditions (Ingram et al., 2004). By definition an endothelial progenitor cell is a cell that can be clonally and serially replated in culture and will give rise to endothelium either by differentiation in vitro or direct incorporation into the vessel wall in vivo. The in vivo method detailed herein is a modification of traditional in vitro collagen gel matrices developed for in vivo implantation following a brief in vitro culture period (Schechner et al., 2000; Enis et al., 2005; Shepherd et al., 2006; Yoder et al., 2007). The method of implanting or injecting extracellular matrices is not new, as evidenced by the many experiments where Matrigel, a decellularized extracellular matrix from a murine tumor, is injected to quantify vascularization ability of the host. As compared to Matrigel implantation, collagen and fibronectin gels fail to recruit substantial host murine vessel ingrowth. Thus, formation of human-murine chimeric vessels is a function of human vascular outgrowth to the host vessels surrounding the implanted gels.

Critical Parameters Isolation, cloning, and propagation of ECFCs Aseptic technique and fresh reagents must be used for all cell culture work. Careful medium changes during the first week of culture are critical for successful culture of ECFCs. Medium must be removed and replaced slowly every 24 hr for the first 7 days, otherwise colony numbers may be diminished. Once ECFC colonies have been established, expansion of cell lines is straightforward. However, ECFC-derived endothelial cell lines have a finite replicative capacity. With continued culture, doubling times will increase and eventually cells will senesce. ECFCs are more frequent in normal UCB than in normal adult peripheral blood. However, there is variability in ECFC frequency among

donors. Disease states and age of the donor may affect the number, time of appearance, population doubling time, or replicative capacity of ECFCs. While culture conditions are optimized for the outgrowth of ECFCs, other cell types are also supported. In some cases, particularly in UCB, MSC colonies will emerge from culture in cEGM-2 (see Fig. 2C.1.1D). Because cEGM-2 contains basic fibroblast growth factor (bFGF), MSC proliferate well in this medium. If MSC colonies arise, it is best to clonally isolate and subculture ECFCs as soon as possible to avoid continued contamination. Phenotypic characterization of ECFCs For assessment of surface antigen expression, keep antibodies and staining samples cold at all times. Protect samples and antibodies from prolonged exposure to light. For best results, process fixed samples on a flow cytometer within 7 days of staining. Fluorescently labeled AcLDL reagents should be used within 1 month of purchase. Counterstaining nuclei with DAPI or other nuclear dye will assist in identification and assessment of the percentage of cells which ingested AcLDL under fluorescent microscopy. Uptake of fluorescently labeled AcLDL can also be assessed using a flow cytometer. Matrigel-coated wells for tube formation assays must be prepared immediately prior to seeding the cells. Do not allow the thin Matrigel coating to dry out. Transplantation of ECFCs into mice Attentive ECFC cell culture is critical for successful vascularization of gel implants. Specifically, ECFCs of low passage, which have been maintained in subconfluent culture conditions, tend to yield better vasculature formation. Continued passaging and maturation of ECFCs correlates to lower vascularization in vivo (Melero-Martin et al., 2007). In the authors’ experience, fresh (i.e., never previously cryopreserved) ECFCs also yield better vascularization. Accurate pH of the gel solution is necessary to cast a cellularized gel with the proper consistency. A high pH tends to yield large, soft gels, which are difficult to handle when implanting. A low pH can produce a small, contracted gel. While pH is controlled in commercially available collagen type I solutions, there remains some lot variability. Performing a trial run of casting a gel without cells may be useful in determining the volume of NaOH

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necessary to attain the proper pH with each new lot of collagen type I. Some experimenters require in vitro tube formation prior to implantation to increase the likelihood of vasculature formation. Experimenters should correlate the extent of in vitro tube formation with formed vasculature to determine if this is a reasonable assertion. The formation of a capillary-like network is vital for formation of vasculature in vivo. A number of immuno-compromised mouse strains can be used as a recipient for the gel implant xenograft model. Studies have been reported using severe combined immunodeficient (SCID)/beige (Schechner et al., 2000; Enis et al., 2005; Shepherd et al., 2006) and nonobese diabetic (NOD)/SCID mice (Yoder et al., 2007). Functional vasculature can be seen within grafts as early as 2 days post-implant. Gel implants are typically harvested between day 2 and 60 (Schechner et al., 2000; Enis et al., 2005; Shepherd et al., 2006; Yoder et al., 2007). Mouse vasculature is not typically found within the border of the gel implant, although extension of host vasculature into the gel is sometimes seen. Thus it is important to ensure that sections are stained with both anti–mouse CD31 and anti–human CD31 to determine the origin of the vasculature. The choice of CD31 is made due to the availability of speciesspecific antibodies suitable for immunohistochemistry of paraffin sections.

See Table 2C.1.2 for information about dealing with problems encountered in these assays.

Transplantation of ECFCs into mice Vascularization of the collagen gel can be seen immediately upon opening the skin. Gel explants range in color from white, indicating very little vasculature formation, to pink or deep red. Gels cast without ADSCs or other supporting cell types are prone to microaneurysms and thus may contain blood clots which will make it difficult to quantify the number of blood vessels. CD31+ vessel density in the ECFC and co-culture implants of 26.6 ± 5.8 and 122.4 ± 9.8 vessels per mm2 , respectively have been reported (K. March, pers. comm). ECFCs alone will typically form vasculature in ∼30% of implanted gels. Additionally, the gels will not typically be vascularized uniformly, with regions of copious vascularity and regions of avascularity. These regions can often be visualized with moderate magnification (such as with a dissecting microscope), allowing the visualization of individual small vessels. Therefore, care must be taken when quantifying the extent of vasculature from a narrow span of the total gel area not to overstate or understate the extent of de novo vasculogenesis. The human vasculature that is formed, in absence of a supporting cell type such as ADSCs, is prone to microaneurysms, is nonuniform in vessel diameter distribution, and lacks a smooth muscle layer.

Anticipated Results

Time Considerations

Initiation and propagation of ECFCs ECFC colonies appear between day 7 and 14 of culture for UCB and between day 14 and 28 for adult peripheral blood. Different disease states may affect the number and time of appearance of ECFC colonies. Adult bloodderived ECFCs can be expanded to 1010 cells after 10 weeks (Hur et al., 2004; Ingram et al., 2004), while UCB-derived ECFCs have higher replicative capacity and can generate as many as 1015 cells after 10 weeks of culture (Ingram et al., 2004). UCB-derived ECFCs have the potential to undergo >50 population doublings (Bompais et al., 2004; Ingram et al., 2004).

Initiation and propagation of ECFCs Isolation of MNCs from peripheral blood or UCB and initial plating of MNC cultures requires ∼3 hr. If processing of UCB cannot be performed immediately, whole anticoagulated UCB can be kept at room temperature with gentle rocking for up to 16 hr. In the authors’ experience, ECFC colonies can be isolated following this holding period; however, the number of colonies will decrease as the time between blood collection and processing increases. Due to the lower frequency of ECFCs, adult peripheral blood samples should be processed immediately after collection.

Troubleshooting

Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

Phenotypic characterization of ECFCs ECFCs uniformly express the endothelial cell–specific surface antigens CD31, CD105, CD144, and CD146, but do not express hematopoietic cell specific surface antigen CD45 or monocyte/macrophage marker CD14. ECFCs will ingest AcLDL and form capillary-like tubes when plated on Matrigel.

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Table 2C.1.2 Troubleshooting Guide for Isolation and Characterization of Endothelial Progenitor Cells Protocols

Problem

Possible cause

Solution

No ECFC colony growth

Harsh treatment of MNCs

Remove and replace medium at a rate of 1 ml/3-4 sec every 24 hr during the first week of culture.

Low MNC seeding density

Seed initial MNC culture at 3-5 × 107 MNCs/well

Reagents are outdated

Use cEGM-2 within 1 month of preparation

Serum is blocking trypsin activity

Wash wells 2-3 times with PBS prior to adding trypsin to remove all prior bound serum

Trypsin activity is low

Use fresh, warm trypsin. Avoid repeated warming and cooling of trypsin.

ECFCs are not dividing

Cells are senescent

All ECFC-derived ECs will eventually senesce. Splitting too severely can lead to premature senescence. Seed cells at 3000-5000 cells/cm2 when subculturing.

ECFC cultures do not express CD31, CD144, or CD146

Cells are not ECFC

If cells do not express CD31, CD144, or CD146, they may be MSCs.

Antibody problem

Test antibodies on known ECs (e.g., HUVECs or HMVECs). Keep antibodies on ice at all times and protect from light.

Cells do not ingest AcLDL

Cells are not endothelial

Use other methods (e.g., analysis of surface antigen expression and Matrigel tube formation) to corroborate phenotype.

Cells do not form tubes when plated on Matrigel

Seeding density is too low

Increase number of cells plated/well.

Suboptimal culture conditions

Use cultures of ECFCs that are 40%-70% confluent for plating on Matrigel.

Cells are not endothelial

Use other methods (e.g., analysis of surface antigen expression and AcLDL uptake) to corroborate phenotype.

ECFCs will not detach

Cellularized Gel pH is too low collagen/fibronectin gels do not solidify

Prior to making cellularized gels with new lots of collagen, ensure proper gel pH by 1N NaOH titer.

Cellularized collagen/fibronectin gels contract in culture

Gel pH is too low

Some contraction is normal, but significant contraction of the gel should be avoided. Prior to making cellularized gels with new lots of collagen, ensure proper gel pH by 1N NaOH titer.

Cellularized collagen/fibronectin gels polymerize, but are large, soft, or fragile

Gel pH is too high

Prior to making cellularized gels with new lots of collagen, ensure proper gel pH by 1N NaOH titer.

Harvested gels lack vasculature

Cell handling

Ensure proper cell handling (see Critical Parameters). Ensure cells have formed tubes in vitro prior to implantation.

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ECFC-derived colonies arise in culture of UCB MNCs between day 5 and day 14. Colonies from one donor can be clonally isolated and serially expanded to multiple 75-cm2 flasks by 4 weeks of culture. Adult peripheral blood ECFCs arise in culture between day 14 and day 28, and can be expanded to a 75-cm2 flask by the sixth week. There is variability in number of primary ECFC colonies and population doubling times among donors. Phenotypic characterization of ECFCs Staining for cell surface antigen expression will take ∼2 hr. Data collection and analysis can be completed in ∼1 hr. Cells can be fixed and stored for up to 1 week if data collection cannot be performed immediately following staining. Assessment of AcLDL ingestion requires preparation of ECFC cultures at least 1 day prior to the assay. Incubation with AcLDL and visualization of ingestion is completed in 5 hr. Matrigel tube-forming assays require 1 hr for set up and 8 to 24 hr for incubation. While all three characterization techniques should be performed to confirm an endothelial phenotype, it is not necessary that they all be performed on the same day. Transplantation of ECFCs into mice With experience, casting of gel implants can be completed within 1 hr. The surgical procedure will take 2 to 4 hr, depending on the number of mice to receive implants. Gel implants can be harvested between day 2 and 60 after implantation.

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Isolation and Characterization of Endothelial Progenitor Cells from Human Blood

properties compared with mature vessel wall endothelial cells. Blood 103:2577-2584. Dimmeler, S., Aicher, A., Vasa, M., MildnerRihm, C., Adler, K., Tiemann, M., Rutten, H., Fichtlscherer, S., Martin, H., and Zeiher, M. 2001. HMG-CoA reductase inhibitors (statins) increase endothelial progenitor cells via the PI 3-kinase/Akt pathway. J. Clin. Invest. 108:391397. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Enis, D.R., Shepherd, B.R., Wang, R., Qasim, A., Shanahan, C.M., Weissberg, P.L., Kashgarian, M., Pober, J.S., and Schechner, J.S. 2005. Induction, differentiation, and remodeling of blood vessels after transplantation of Bcl-2-transduced endothelial cells. Proc. Natl. Acad. Sci. U.S.A 102:425-430. Gulati, R., Jevremovic, D., Peterson, T.E., Chaterjee, S., Shah, V., Vile, R.G., and Simon, R.D. 2003. Diverse origin and function of cells with endothelial phenotype obtained from adult human blood. Circ. Res. 93:1023-1025. Hassan, N.F., Campbell, D.E., and Douglas, S.D. 1986. Purification of human monocytes on gelatin-coated surfaces. J. Immunol. Methods 95:273-276. Hill, J.M., Zalos, G., Halcox, J.P., Schenke, W.H., Waclawiw, M.A., Quyyumi, A.A., and Finkel, T. 2003. Circulating endothelial progenitor cells, vascular function, and cardiovascular risk. N. Engl. J. Med. 348:593-600. Hur, J., Yoon, C.H., Kim, H.S., Choi, J.H., Kang, H.J., Hwang, K.K., Oh, B.H., Lee, M.M., and Park, Y.B. 2004. Characterization of two types of endothelial progenitor cells and their different contributions to neovasculogenesis. Arterioscler. Thromb. Vasc. Biol. 24:288-293. Ingram, D.A., Mead, L.E., Tanaka, H., Meade, V., Fenoglio, A., Mortell, K., Pollok, K., Ferkowicz, M.J., Gilley, D., and Yoder, M.C. 2004. Identification of a novel hierarchy of endothelial progenitor cells utilizing human peripheral and umbilical cord blood. Blood 104:2752-2760. Ito, H., Rovira, I.I., Bloom, M.L., Takeda, K., Ferrans, V.J., Quyyumi, A.A., and Finkel, T. 1999. Endothelial progenitor cells as putative targets for angiostatin. Cancer Res. 59:58755877.

Bancroft, J.D. and Gamble, M. 2002. Theory and Practice of Histological Techniques. 5th Ed.. Churchill Livingstone, New York.

Kalka, C., Masuda, H., Takahashi, T., Kalka-Moll, W.M., Silver, M., Kearney, M., Li, T., Isner, J.M., and Asahara, T. 2000. Transplantation of ex vivo expanded endothelial progenitor cells for therapeutic neovascularization. Proc. Natl. Acad. Sci. U.S.A. 97:3422-3427.

Baumgarth, N. and Roederer, M. 2000. A practical approach to multicolor flow cytometry for immunophenotyping. J. Immunol. Methods 243:77-97.

Lin, Y., Weisdorf, D.J., Solovey, A., and Hebbel, R.P. 2000. Origins of circulating endothelial cells and endothelial outgrowth from blood. J. Clin. Invest. 105:71-77.

Bompais, H., Chagraoui, J., Canron, X., Crisen, M., Liu, X.H., Anjo, A., Tolla-LePort, C., Leboef, M., Charbord, P., Bikfalvi, A., and Uzan, G. 2004. Human endothelial cells derived from circulating progenitors display specific functional

Melero-Martin, J.M., Khan, Z.A., Picard, A., Wu, X., Paruchuri, S., and Bischoff, J. 2007. In vivo vasculogenic potential of human blood-derived endothelial progenitor cells. Blood 109:47614768.

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Perfetto, S.P., Ambrozak, D., Nguyen, R., Chattopadhyay, P., and Roederer, M. 2006. Quality assurance for polychromatic flow cytometry. Nat. Protoc. 1:1522-1530. Prater, D.N., Case, J., Ingram, D.A., and Yoder, M.C. 2007. Working hypothesis to redefine endothelial progenitor cells. Leukemia 21:11411149. Rafii, S. and Lyden, D. 2003. Therapeutic stem and progenitor cell transplantation for organ vascularization and regeneration. Nat. Med. 9:702712. Rehman, J., Li, J., Orschell, C.M., and March, K.L. 2003. Peripheral blood “endothelial progenitor cells” are derived from monocyte/macrophages and secrete angiogenic growth factors.” Circulation 107:1164-1169. Schechner, J.S., Nath, A.K., Zheng, L., Kluger, M.S., Hughes, C.C., Sierra-Honigmann, M.R., Lorber, M.I., Tellides, G., Kashgarian, M., Bothwell, A.L., and Pober, J.S. 2000. In vivo formation of complex microvessels lined by human endothelial cells in an mmunodeficient mouse. Proc. Natl. Acad. Sci. U.S.A. 97:9191-9196. Schmeisser, A., Graffy, C., Daniel, W.G., and Strasser, R.H. 2003. Phenotypic overlap be-

tween monocytes and vascular endothelial cells. Adv. Exp. Med. Biol. 522:59-74. Shapiro, H.M. 2003. Practical Flow Cytometry. 4th Edition. Wiley-Liss, Wilmington, Del. Shepherd, B.R., Enis, D.R., Wang, F., Suarez, Y., Pober, J.S., and Schechner, J.S. 2006. Vascularization and engraftment of a human skin substitute using circulating progenitor cell-derived endothelial cells. Faseb J. 20:1739-1741. Werner, N., Kosiol, S., Schiegl, T., Ahlers, P., Walenta, K., Link, A., B¨ohm, M., and Nickenig, G. 2005. Circulating endothelial progenitor cells and cardiovascular outcomes. N. Engl. J. Med. 353:999-1007. Yoder, M.C., Mead, L.E., Prater, D., Krier, T.R., Mroueh, K.N., Li, F., Krasich, R., Temm, C.J., Prchal, J.T., and Ingram, D.A. 2007. Redefining endothelial progenitor cells via clonal analysis and hematopoietic stem/progenitor cell principals. Blood 109:1801-1809. Ziegelhoeffer, T., Fernandez, B., Kostin, S., Heil, M., Voswinckel, R., Helisch, A., Kostin S, Heil M, Voswinckel R., and Helisch, A. 2004. Bone marrow-derived cells do not incorporate into the adult growing vasculature. Circ. Res. 94:230238.

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2C.1.27 Current Protocols in Stem Cell Biology

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Derivation of Epicardium-Derived Progenitor Cells (EPDCs) from Adult Epicardium

UNIT 2C.2

Nicola Smart1 and Paul R. Riley1 1

UCL Institute of Child Health, London, United Kingdom

ABSTRACT The epicardium has, like the other cell lineages of the terminally differentiated adult heart, long been regarded as quiescent, incapable of migration or differentiation. In contrast, the embryonic epicardium possesses an innate ability to proliferate, migrate, and differentiate into a number of mature cardiovascular cell types, including vascular smooth muscle cells, fibroblasts, cardiomyocytes, and, arguably, some endothelial cells. In recapitulating its essential developmental role, we recognized the ability of the actin-binding peptide thymosin β4 (Tβ4) to induce epicardium-derived progenitor cell (EPDC) migration from adult heart and noted the derivation of cell types originating from embryonic epicardium. This protocol provides a means of enabling adult EPDC outgrowth and culture. We establish a model system in which to study the ability of factors to influence the migration of vascular precursors and their differentiation and to move towards screening of small molecules ex vivo prior to clinical trials of therapeutic cardiac repair. Curr. Protoc. Stem C 2009 by John Wiley & Sons, Inc. Cell Biol. 8:2C.2.1-2C.2.9.  Keywords: epicardium r adult EPDCs r thymosin β4 r adult heart

INTRODUCTION This unit includes a protocol for the outgrowth and culture of epicardium-derived cells (EPDCs) from the adult epicardium. In the developing embryo, the epicardium is the principal source of precursor cells for coronary vasculogenesis (Perez-Pomares et al., 2006). More recently, the epicardium has been shown to contribute ∼4% of the cardiomyocytes of the fully developed heart (Cai et al., 2008; Zhou et al., 2008). Embryonic EPDCs possess an innate capacity for migration and are thus readily isolated and cultured (Chen et al., 2002). However, this capacity rapidly diminishes over the course of development and is virtually lost by adulthood. Having identified thymosin β4 (Tβ4) as a peptide that is required for embryonic EPDC migration and coronary vasculature formation, the authors of this unit demonstrated that this factor could indeed induce EPDC migration from adult heart (Smart et al., 2007). Protocols exist for the derivation of EPDCs from adult hearts (van Tuyn et al., 2006); however, unstimulated adult EPDCs emerge only very slowly (4 to 7 days) and, in our hands, readily differentiate and are therefore difficult to isolate as progenitor cells. The addition of Tβ4 in this protocol stimulates extensive migration of proliferating EPDCs, enabling their derivation within 24 to 48 hr, prior to any differentiation event and with a considerably higher yield. This unit describes the basic method for derivation of EPDCs in tissue culture dishes (and, optionally, on coverslips for subsequent immunofluorescence analysis) in the Basic Protocol. EPDCs remain largely as undifferentiated progenitor cells for the initial 24 to 48 hr post-outgrowth, but, under the conditions employed, the majority spontaneously differentiate over the subsequent 2 to 4 days. A Support Protocol is provided Somatic Stem Cells Current Protocols in Stem Cell Biology 2C.2.1-2C.2.9 Published online February 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02c02s8 C 2009 John Wiley & Sons, Inc. Copyright 

2C.2.1 Supplement 8

Figure 2C.2.1 Derivation of epicardium-derived cells (EPDCs) as cardiovascular progenitors from the adult heart. Thymosin β4 (Tβ4) stimulates outgrowth of large colonies of EPDCs from adult heart explants (A). Gata5-EYFP lineage trace analysis (B) and epicardin expression (C) confirm the epicardial origin of outgrowing cells. Following migration, EPDCs differentiate into vasculogenic cells including smooth muscle cells (D), fibroblasts (E), and endothelial cells (F), as well as cardiac progenitors which are positive for Nkx2.5 (G) and Isl-1 (H). EPDCs proliferate upon migration from adult explants (Ki67, as shown in G, H).

that details the use of tested antibodies for immunostaining to assess the differentiation status of EPDCs and identify the cell types produced in the cultures following differentiation. Optionally, if genetic lineage tracing of epicardial cells is desired, explant cultures can be prepared using hearts from a suitable mouse line, such as a Gata5Cre × R26R-EYFP cross which will label epicardial derivatives with EYFP fluorescence (Fig. 2C.2.1B). Derivation of EpicardiumDerived Progenitor Cells (EPDCs) from Adult Epicardium

NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to government regulations for the care and use of laboratory animals.

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Current Protocols in Stem Cell Biology

THYMOSIN β 4–INDUCED OUTGROWTH OF ADULT EPICARDIUM-DERIVED CELLS (EPDCs)

BASIC PROTOCOL

This protocol describes the steps for isolating EPDCs from adult mouse heart. The authors have found that thymosin β4 stimulates the outgrowth and migration of the EPDCs. NOTE: All tissue culture reagents and materials must be sterile. Dissection tools should ideally be autoclaved or, alternatively, placed in 70% ethanol for 5 min and air dried inside the tissue culture hood before use. NOTE: All tissue preparation steps are performed in a laminar flow hood and, if desired, under a stereomicroscope.

Materials 0.1% gelatin solution (see recipe) 8- to 12-week-old adult mice (C57Bl/6 strain used; other strains and ages not tested) Dulbecco’s phosphate-buffered saline (DPBS; Invitrogen, cat. no. 14190) EPDC culture medium (see recipe) supplemented with 100 ng/ml thymosin β4 (see recipe) Tissue culture dishes or plates of desired size for culture (Table 2C.2.1) and (optionally) glass coverslips of the appropriate size (also in Table 2C.2.1) Forceps (0.5-mm approximate tip size), sterile Dissection scissors, sterile Sterile 60- or 100-mm culture/bacteriological dish (not gelatin coated) for dissection Scalpel blade Humidified 37◦ C, 5% CO2 incubator Additional reagents and equipment for sacrifice of mice by cervical dislocation (Donovan and Brown, 2006) Dissect heart 1. Coat culture dishes, plates, or coverslips with gelatin by pipetting the appropriate volume of 0.1% gelatin solution (see “culture volume” column in Table 2C.2.1) per dish or plate well and allowing to stand for 15 min. Aspirate the gelatin solution. It is advisable to culture EPDCs on coverslips if cells are to be analyzed by immunofluorescence (Support Protocol). In this case, coverslips should be placed into the culture dish prior to gelatin coating. If desired, an additional non-gelatin-coated coverslip may be placed over the tissue pieces to reduce tissue floating and encourage adhesion (step 10, below).

2. Sacrifice adult mouse by cervical dislocation (Donovan and Brown, 2006). Table 2C.2.1 Recommended Parameters for EPDC Culture in Various Plate Formats

Culture volume (ml)a

Coverslip size (mm)

Amount tissue/well

12-well plate

0.8

13

1/8 heart

6-well plate

2.0

18

1/4 heart

35-mm dish

2.0

18

1/4 heart

60-mm dish

4.0

Not recommended

1/2 heart

100-mm dish

10.0

Not recommended

1 to 2 hearts

TC plate format

a Volumes given apply to each well of multiwall plates.

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3. Using sterile forceps and scissors, make a lateral incision in the center of the abdomen and tear back the fur to expose the rib cage. 4. Carefully cut upwards through the sternum and along the diaphragm, taking care not to cut into the heart. Pull back the ribs to reveal the heart. 5. Remove the heart using forceps and dissect away the major vessels. 6. Place tissue in a 60-mm tissue culture dish (not gelatin coated) containing 2 ml DPBS. Cut the heart into quarters and allow blood to rinse from the tissue. Carefully aspirate away DPBS. Using a sterile scalpel, mince the heart into pieces of ∼0.5 to 1 mm3 . IMPORTANT NOTE: Reproducible EPDC outgrowth strongly depends upon the size of the heart pieces (optimally 0.5 to 1 mm3 ). Larger pieces will not adhere to permit sufficient migration, while smaller pieces tend to dissociate completely, and cardiomyocyte death precedes adherence and EPDC outgrowth.

Seed fragments for outgrowth 7. Divide the heart pieces into equal portions of the appropriate size (for example, one adult heart is typically divided between four wells of a 6-well plate for optimal EPDC outgrowth; refer to Table 2C.2.1 for other dish sizes). 8. Pipet the appropriate volume of EPDC culture medium containing 100 ng/ml thymosin β4 into each dish or plate well to be used (for recommended volumes, refer to Table 2C.2.1). 9. Place one portion of heart tissue into the center of each dish or plate well and ensure that all pieces are submerged. 10. Optional: Carefully place a round glass coverslip (not gelatin coated) over the heart pieces to prevent the tissue from floating. 11. Gently transfer the plate to a humidified 37◦ C, 5% CO2 incubator. Maintain cultures with minimum disturbance to allow explants to adhere. No feeding is required for the first 48 hr. IMPORTANT NOTE: Minimal disturbance is absolutely essential for EPDC outgrowth. Explants adhere only tenuously at first, and disturbance in the earliest days of culture will prevent adhesion or lead to detachment. Plates should be transferred extremely cautiously between incubator and microscope or culture hood. After sufficient EPDCs have emerged, explants attach more firmly, but care is still required as detachment may easily occur.

Culture EPDCs 12. After 24 to 48 hr in culture, transfer the plate to the culture hood, taking great care to avoid disturbing the explants. Pipet an appropriate volume of DPBS (see “culture volume” column in Table 2C.2.1) slowly into the dish, directing the solution toward the rim of the plate and not directly at the explant. Aspirate DPBS and repeat this process for a second wash. 13. Add 2 ml EPDC medium, freshly supplemented with 100 ng/ml thymosin β4. EPDCs can be harvested as progenitor cells at this stage or left for a further 2 to 4 days for differentiation to occur, prior to assessment of cellular phenotype. Derivation of EpicardiumDerived Progenitor Cells (EPDCs) from Adult Epicardium

Following their emergence from the explant at 24 to 48 hr, EPDCs display a “cobblestone” morphology, characteristic of epithelial cells (Fig. 2C.2.1, panel A). Following migration, cells at the outer edges of the explant, at least when cultured under the conditions described herein, spontaneously differentiate into a number of discernable cell types, including smooth muscle cells, fibroblasts, endothelial cells, and cardiomyocyte progenitors (for phase-contrast images of cellular morphology, please refer to Smart et al., 2007; differentiated cell types are shown in Fig. 2C.2.1, panels D-H).

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Current Protocols in Stem Cell Biology

CHARACTERIZATION OF EPDC PHENOTYPES BY IMMUNOFLUORESCENCE

SUPPORT PROTOCOL

Following their migration and expansion in culture, EPDCs can be utilized in a range of experimental settings that will ultimately require an assessment of the different cell types derived following differentiation. Embryonic EPDCs have been characterized as progenitors, and following differentiation, by RT-PCR (Chen et al., 2002). This approach could be utilized with adult EPDCs to identify the range of cell types within a single culture. Alternatively, this support protocol may be used for an immunological assessment of individual EPDCs cultured on glass coverslips. The protocol provided below details antibodies that have been successfully utilized to characterize undifferentiated EPDCS and their derivatives, including cardiac progenitors, primitive cardiomyocytes, vascular smooth muscle cells, endothelial cells, and fibroblasts (Table 2C.2.2). See Table 2C.2.1 for appropriate volumes.

Materials Heart explants cultured on glass coverslips (Basic Protocol) 4% (w/v) paraformaldehyde in PBS (freshly prepared) Phosphate-buffered saline (PBS; prepared according to manufacturer’s instructions from PBS tablets; Sigma, cat. no. P-4417) Blocking solution containing 0.1% (v/v) Triton X-100 Blocking solution (see recipe) Primary antibodies of choice (refer to Table 2C.2.2) Appropriate fluorochrome-conjugated secondary antibody (against Ig of species in which primary antibody was raised) 5 μg/ml Hoechst 33342 in PBS Suitable commercially available mounting medium or 50% (v/v) glycerol in PBS Microscope slides Fluorescence microscope with appropriate filters for fluorochrome used Table 2C.2.2 Antibody Sources and Conditions for Immunofluorescence-Based Characterization of EPDCs

Antibody

Supplier

Clonality

Source

Dilution

Epicardin (TCF21)

Abcam

Polyclonal

Rabbit

1:100

WT-1

Abcam

Monoclonal

Rabbit

1:50

TBX18

Chemicon

Monoclonal

Mouse

1:50

GATA-5

Abcam

Polyclonal

Rabbit

1:100

Ki67

Dako

Monoclonal

Rat

1:30

ISL-1

Developmental Studies Hybridoma Bank

Monoclonal

Mouse

1:30

NKX2.5

Santa Cruz

Polyclonal

Rabbit

1:50

GATA-4

Santa Cruz

Polyclonal

Rabbit

1:50

α-Sarcomeric actinin

Sigma

Monoclonal

Mouse

1:500

cTNT

Abcam

Polyclonal

Rabbit

1:200

Procollagen type I

Santa Cruz

Polyclonal

Goat

1:100

α-Smooth muscle actin

Sigma

Monoclonal

Mouse

1:500

Flk1

BD Pharmingen

Monoclonal

Rat

1:50 Somatic Stem Cells

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Fix cells 1. Culture adult heart explants on glass coverslips as described (Basic Protocol). 2. After 24 hr to 5 days of culture, fix cells with 4% paraformaldehyde for 10 min at room temperature. EPDCs typically emerge from the explant after 24 to 48 hr in culture. At these early time points, EPDCs exist largely as undifferentiated progenitors. Subsequently (days 3 to 5), the majority of EPDCs spontaneously differentiate, as described above. Cultures should be harvested at the appropriate time point, depending on whether the study requires undifferentiated EPDCs (harvest at 24 hr) or differentiated cells (smooth muscle, endothelial cells, and fibroblasts have been detected after 5 days in culture).

3. Pipet an appropriate volume of PBS (see “culture volume” column in Table 2C.2.1) slowly onto the side of the dish, allowing it to gently flow over the cells. Do not pipet forcefully onto the coverslips as this will cause cells to become detached. Leave for 1 min, aspirate PBS, and repeat this process for a second wash.

Permeabilize and block 4. Permeabilize cells with 0.5% Triton X-100 in PBS for 5 min at room temperature. 5. Wash coverslips twice with PBS, as described in step 3. 6. Block nonspecific binding by incubating cells in blocking solution containing 0.1% Triton X-100 for 1 hr at room temperature.

Stain cells with antibody 7. Incubate cells with an appropriate dilution of primary antibody in blocking solution/0.1% Triton X-100, overnight at 4◦ C (for recommended antibody dilutions, refer to Table 2C.2.2). 8. Pipet an appropriate volume of blocking solution/0.1% Triton X-100 (see “culture volume” column in Table 2C.2.1) slowly onto the side of the dish, allowing it to gently flow over the cells. Do not pipet forcefully onto the coverslips as this will cause cells to become detached. Leave for 5 min, aspirate solution, and repeat this process two more times for a total of three washes. Rinse twice in blocking solution (without Triton) using this same technique. 9. Incubate cells with the appropriate secondary antibody diluted according to the manufacturer’s instructions in blocking solution. 10. Pipet an appropriate volume of PBS (see “culture volume” column in Table 2C.2.1) slowly onto the side of the dish, allowing it to gently flow over the cells. Do not pipet forcefully onto the coverslips as this will cause cells to become detached. Leave for 5 min, aspirate PBS, and repeat this process again for a second wash.

Stain nuclei and mount 11. Optional: To stain nuclei, incubate with 5 μg/ml Hoechst in PBS for 5 min at room temperature. 12. Wash cells twice in PBS, as described in step 10. 13. Mount coverslips on microscope slides using mounting medium or 50% glycerol in PBS, and visualize using a fluorescence microscope. Derivation of EpicardiumDerived Progenitor Cells (EPDCs) from Adult Epicardium

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Current Protocols in Stem Cell Biology

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Blocking solution Prepare PBS containing 10% (v/v) normal goat serum and 1% (w/v) bovine serum albumin. EPDC culture medium Supplement DMEM (containing GlutaMax-I and 4.5 g/liter glucose; Invitrogen) with 15% (v/v) fetal bovine serum (FBS), 100 U/ml penicillin, and 100 μg/ml streptomycin. Store at 4◦ C for up to 1 month. Do not supplement with thymosin β4 (see recipe below) until ready to use.

EPDC culture medium containing 100 ng/ml thymosin β4 To prepare 1000× stock (100μg/ml), dilute 1 mg of thymosin β4 (Immunodiagnostik) into 10 ml sterile DPBS. Aliquot and store at −80◦ C until required. Avoid repeated freezing and thawing. When required, dilute 1 μl of the 1000× stock per ml of EPDC culture medium (see recipe) for a final concentration 100 ng/ml, immediately prior to use.

Gelatin solution, 0.1% (w/v) Dissolve 0.5 g of gelatin (from porcine skin) in 500 ml distilled water and autoclave. Store at room temperature for 2 to 3 months.

COMMENTARY Background Information In the adult, the need to maintain both myocardial homeostasis and a healthy coronary vasculature is highlighted by the devastating consequences of coronary artery disease, which frequently results in extensive myocardial necrosis, vessel loss, and subsequent cardiac failure. Resident cardiac progenitor cells have recently been identified (reviewed in Smart and Riley, 2008) which could potentially fulfill the requirements of continued replacement of senescent cells and regeneration of the heart following injury. However, the regenerative capacity of the human myocardium remains inadequate to compensate for the severe loss of heart muscle that follows myocardial infarction. Current research focuses on discovering suitable cell populations for myocardial regeneration and neovascularization and, in parallel, on identifying factors for therapeutic stimulation of resident cardiac progenitor cells to harness their potential for repair. The ability to mobilize endogenous progenitor cells from within the adult heart and to induce their differentiation into cardiomyocytes and vascular cells capable of forming vessels offers tremendous potential for the treatment of human heart disease (Srivastava and Ivey, 2006). In this regard, epicardium-derived cells

represent a therapeutic prospect, subject to the identification of suitable factors to unleash the myogenic and vasculogenic potential of adult epicardium. Primary epicardial cells have been derived from fetal and early neonatal hearts (Chen et al., 2002). Cultures assume an epithelial morphology, express epicardial markers, and can be maintained for at least four passages without alteration in epithelial morphology (Chen et al., 2002). However, the potential of the epicardium, both in terms of its trophic activity in stimulating cardiomyocyte proliferation (Chen et al., 2002) and capacity to migrate (Smart et al., 2007) diminishes rapidly between E12 and postnatal day 4 (P4). The derivation of EPDCs is dependent upon their migration, and this protocol is therefore limited in its application to use with embryonic and neonatal hearts. The precise relationship between EPDC migration and proliferative capacity has not been thoroughly evaluated; however, since EPDCs readily proliferate in culture (Fig. 2C.2.1G,H), it may be that migration away from the explant is sufficient to stimulate EPDC proliferation. Maximal outgrowth is observed at E10.5, a stage in development coincident with the formation of the epicardium; outgrowth diminishes

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considerably by E12.5 and continues to do so such that, by P1, outgrowth is reduced to ∼10% of that at E10.5. In untreated adult explants there is virtually no detectable outgrowth, with only a few isolated cells observed in the culture dish, consistent with the adult epicardium residing in a quiescent state, having lost migration, differentiation, and signaling capacities during the latter half of gestation (Chen et al., 2002). Having demonstrated a requirement for thymosin β4 (Tβ4) in coronary vasculogenesis, angiogenesis, and arteriogenesis in the developing embryo, we investigated its potential to stimulate these same processes in the adult heart (Smart et al., 2007). In contrast to untreated adult heart, Tβ4 stimulated extensive outgrowth of cells (Fig. 2C.2.1, panel A) which, like those obtained in embryonic cultures, display a characteristic epithelial morphology and are positive for the epicardial-specific transcription factor, epicardin/TCF21, as well as proteins associated with the active embryonic epicardium, such as WT-1, TBX18, and GATA-5. Following migration away from the explant, EPDCs proliferate (Ki67 positive) and differentiate into a variety of discernable cell types, known to derive from the embryonic epicardium. Cardiac progenitors are detected by virtue of their coexpression of ISL-1, NKX2.5, and GATA-4 (at 24 to 48 hr of culture; Fig. 2C.2.1G,H). Following removal of the explant and a further 3 days in culture (day 5), large differentiated cells are detected which weakly express α-sarcomeric actinin, cardiac troponin T, and cardiac myosin-binding protein C. However, under the culture conditions employed, no mature, fully differentiated cardiomyocytes with definitive sarcomeric structure are observed. Procollagen type I, α-smooth muscle actin, and Flk1 positive cells indicate the presence of fibroblasts, smooth muscle, and a limited number of endothelial cells, respectively (Fig. 2C.2.1, panels D to F). Thus, Tβ4induced adult EPDCs represent a viable source of therapeutic cardiomyogenic and vascular progenitors.

Critical Parameters and Troubleshooting

Derivation of EpicardiumDerived Progenitor Cells (EPDCs) from Adult Epicardium

In our hands, the degree of EPDC outgrowth can be extremely variable, but strongly depends upon the following critical factors. Size of heart pieces EPDC outgrowth depends upon the size of the heart pieces, which should optimally be between 0.5 and 1 mm3 . Larger pieces will not

adhere to permit sufficient migration, while smaller pieces tend to dissociate completely and cardiomyocyte death precedes adherence and EPDC outgrowth. Minimal disturbance of explants prior to outgrowth Minimal disturbance is absolutely essential for EPDC outgrowth. Explants adhere only tenuously in the first instance, and disturbance in the first days of culture will prevent adhesion or lead to detachment. Plates should be transferred extremely carefully between the incubator and microscope or culture hood. After sufficient EPDCs have emerged, explants attach more firmly, but care is still required as detachment may easily occur. Activity of Tβ4 We have experienced considerable variability between batches of Tβ4, which profoundly affects the degree of EPDC outgrowth. We are not aware of any simple assay for the biological activity of Tβ4, but it may be desirable to confirm the reported activation of signaling mechanisms, as reported for the Akt pathway in C2C12 myoblasts (Bock-Marquette et al., 2004; Smart et al., 2007). Other sources of Tβ4 are now commercially available (Abcam, ProSpecBio) but these have not been tested for EPDC outgrowth.

Anticipated Results This protocol uses Tβ4 to stimulate “quiescent” adult EPDCs, enabling their migration and subsequent differentiation. The degree of outgrowth from adult heart explants varies considerably. Not all heart pieces in a single preparation produce outgrowths, but those that do typically yield 30 to 3000 EPDCs after 48 hr. The method may be applied to the study of other putative angiogenic or cardiomyogenic factors, either alone or in combination with Tβ4, to assess regenerative potential. In this context, we found that VEGF, FGFs, and AcSDKP led to a significant increase in the numbers of Tie2/Flk1 positive endothelial cells derived from both embryonic and adult EPDC cultures (Smart et al., 2007). Extending this protocol to assess EPDC migration from mutant adult mouse hearts should provide valuable insight into the epicardial lineage per se, the mechanisms underlying (coronary) vasculogenesis, and cellular commitment toward formation of de novo myocardium. From a translational standpoint, models such as this will be invaluable for screening of small molecules for drug

2C.2.8 Supplement 8

Current Protocols in Stem Cell Biology

discovery prior to clinical trials of therapeutic cardiac repair.

Time Considerations This protocol is both relatively straightforward and rapid in terms of hands-on time. Preparation of the tissue for EPDC culture can be achieved in <1 hr. Depending on the experimental study, cultures must then be left for 1 to 5 days for appropriate outgrowth and/or differentiation. The Support Protocol is a typical immunofluorescence protocol and takes 16 to 24 hr to complete.

Acknowledgements Our research is funded by the British Heart Foundation and the Medical Research Council.

Literature Cited Bock-Marquette, I., Saxena, A., White, M.D., Dimaio, J.M., and Srivastava, D. 2004. Thymosin β4 activates integrin-linked kinase and promotes cardiac cell migration, survival and cardiac repair. Nature 432:466-472. Cai, C.L., Martin, J.C., Sun, Y., Cui, L., Wang, L., Ouyang, K., Yang, L., Bu, L., Liang, X., Zhang, X., Stallcup, W.B., Denton, C.P., McCulloch, A., Chen, J., and Evans, S.M. 2008. A myocardial lineage derives from Tbx18 epicardial cells. Nature 454:104-108. Chen, T.H., Chang, T.C., Kang, J.O., Choudhary, B., Makita, T., Tran, C.M., Burch, J.B.E., Eid, H., and Sucov, H.M. 2002. Epicardial induction of fetal cardiomyocyte proliferation via

a retinoic acid-inducible trophic factor. Devel. Biol. 250:198-207. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Perez-Pomares, J.M., Mironov, V., Guadix, J., Macias, D., Markwald, R., and Munoz-Chapuli, R. 2006. In vitro self-assembly of proepicardial cell aggregates: An embryonic vasculogenic model for vascular tissue engineering. Anat. Rec. Part A 288A:700-713. Smart, N. and Riley, P.R. 2008. The stem cell movement. Circ. Res. 102:1155-1168. Smart, N., Risebro, C.A., Melville, A.A., Moses, K., Schwartz, R.J., Chien, K.R., and Riley, P.R. 2007. Thymosin beta-4 is essential for coronary vessel development and promotes neovascularization via adult epicardium. Ann. N.Y. Acad. Sci. 1112:171-188. Srivastava, D. and Ivey, K.N. 2006. Potential of stem-cell-based therapies for heart disease. Nature 441:1097-1099. van Tuyn, J., Atsma, D.E., Winter, E.M., van der Velde-van Dijke, I., Pijnappels, D.A., Bax, N.A.M., Knaan-Shanzer, S., Gittenberger–de Groot, A.C., Poelmann, R.E., van der Laarse, A., van der Wall, E.E., Schalij, M.J., and de Vries, A.A. 2006. Epicardial cells of human adults can undergo an epithelial-to-mesenchymal transition and obtain characteristics of smooth muscle cells in vitro. Stem Cells 2006-0366. Zhou, B., Ma, Q., Rajagopal, S., Wu, S.M., Domian, I., Rivera-Feliciano, J., Jiang, D., von, G.A., Ikeda, S., Chien, K.R., and Pu, W.T. 2008. Epicardial progenitors contribute to the cardiomyocyte lineage in the developing heart. Nature 454:109-113.

Somatic Stem Cells

2C.2.9 Current Protocols in Stem Cell Biology

Supplement 8

Isolation and Expansion of Cardiosphere-Derived Stem Cells

UNIT 2C.3

Jun-Jie Tan,1 Carolyn A. Carr,1 Daniel J. Stuckey,1 Georgina M. Ellison,2 Elisa Messina,3 Alessandro Giacomello,3 and Kieran Clarke1 1

Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom 2 Stem Cell and Molecular Physiology Laboratory, Research Institute for Sport and Exercise Sciences, Liverpool John Moores University, Liverpool, United Kingdom 3 Department of Experimental Medicine, Cenci-Bolognetti Foundation, Pasteur Institute, University of Rome ‘Sapienza,’ Rome, Italy

ABSTRACT The isolation and in vitro expansion of stem cells from the adult heart may provide a cell therapy for regenerating damaged myocardium. Cardiac stem cells can be isolated via magnetic or fluorescent cell sorting using specific cell-surface markers, including c-kit, Sca-1, and Isl-1. Because these isolation methods yield relatively few cells, substantial in vitro expansion is required to generate sufficient cell numbers for therapy. An alternative method uses cells spontaneously shed from cultured heart explants, which are harvested, induced to form cardiospheres, and expanded as a monolayer for several passages. This method for generating therapeutically relevant numbers of cells in a shorter time period than cell surface marker–based isolations is ideally suited for autologous cardiac stem cell therapy after myocardial infarction. Curr. Protoc. Stem Cell Biol. 16:2C.3.1-2C.3.12. C 2011 by John Wiley & Sons, Inc.  Keywords: cardiac stem cells r cardiospheres r cardiosphere-derived cells r myocardial regeneration therapy

INTRODUCTION Multipotent cardiac stem cells, able to undergo differentiation into cardiomyocytes, smooth muscle, and endothelial cells, have been identified in the adult heart using cellsurface markers (Beltrami et al., 2003; Oh et al., 2003; Tateishi et al., 2007). This unit adapts a technique for isolating and expanding cardiac stem cells, originally described by Messina et al. (2004) and Smith et al. (2007), with modifications relevant for human biopsies (Messina et al., 2004; Smith et al., 2007; Davis et al., 2010). This protocol begins with excision of a neonatal rat heart, which is then minced and explanted onto a petri dish to allow cells to migrate from the explants. These explant–derived cells are cultured for 7 days to reach 80% to 90% confluency, and are selectively harvested using their cardiosphere-forming capability, which can be induced using cardiosphere growth medium (CGM). Cardiospheres are then cultured and expanded as a monolayer. Cardiosphere-derived cells (CDC) can be maintained in vitro, have stem cell properties, and retain the ability to re-form cardiospheres. The whole procedure can be completed within 16 days. NOTE: All procedures are performed in the sterile cell culture Class II cabinet. NOTE: All solutions are prepared in a sterile environment and kept according to manufacturer’s instruction. NOTE: All explants, cardiospheres, and cardiosphere-derived cell cultures are kept in a humidified incubator at 37◦ C with 5% CO2 . Current Protocols in Stem Cell Biology 2C.3.1-2C.3.12 Published online February 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc02c03s16 C 2011 John Wiley & Sons, Inc. Copyright 

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BASIC PROTOCOL 1

NEONATAL RAT HEART EXCISION, PROCESSING, AND EXPLANT CULTURE This protocol describes the excision, processing and culture of heart explants. Explants can be maintained in complete explant medium (CEM). Explant-derived cells can be isolated every 6 to 7 days by replating the explants, up to 4 times. NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to government regulations for the care and use of laboratory animals.

Materials 1 mg/ml fibronectin stock solution from bovine plasma (Sigma) Dulbecco’s phosphate-buffered saline without CaCl2 and MgCl2 (DPBS, Sigma) Complete explant medium (CEM, see recipe) 0.05% trypsin/EDTA (Invitrogen), 37◦ C Sprague-Dawley neonatal rat 60-mm sterile tissue culture–treated petri dishes Dumont no.7 forceps, sterilized Fine iris scissors, sterilized Inverted light microscope Prepare dishes for explant culture 1. Coat eight 60-mm sterile petri dishes with fibronectin as follows: a. Dilute 1 mg/ml fibronectin (from bovine plasma) to 4 μg/ml in DPBS without CaCl2 and MgCl2 . b. Add 1 ml of this solution per 60-mm cell culture–treated petri dish. c. Leave the petri dish at 37◦ C for 30 min. d. Remove the solution and wash once with DPBS. Do not let the fibronectin-coated dish dry out. Care should be taken not to scratch the fibronectin coating when removing the solution/washings.

2. Add 1.5 ml of CEM to each fibronectin-coated petri dish and replace in the incubator. The fibronectin-coated dishes are to be used for the explant cultures. The dishes containing the 1.5 ml of CEM can be kept in the incubator until use, but we recommend that the coated dishes be used on the day of preparation.

3. Prior to heart isolation, prepare two uncoated petri dishes filled with 5 ml sterile DPBS (for tissue processing), one filled with 2 ml of prewarmed 0.05% trypsin (for trypsin digestion), and one filled with 2 ml of CEM (for trypsin inactivation).

Isolate neonatal rat heart and prepare explant culture 4. Sacrifice the neonatal rat by cervical dislocation, rapidly excise the heart using sterile scissors, and place it in the one of the petri dishes with DPBS prepared in step 3. 5. Cut the heart into several pieces and rinse with DPBS to remove blood. Using the sterile forceps, transfer the heart tissue to the second petri dish containing DPBS for further washing. Isolation and Expansion of CardiosphereDerived Stem Cells

Multiple washes of the heart tissue with DPBS are necessary because excessive blood may affect cell growth. Alternatively, the whole heart may be perfused with DPBS.

6. Transfer the heart tissue to the petri dish prepared in step 3 containing trypsin and keep in the solution for 3 min while mincing the tissue into several pieces using sterile fine iris scissors.

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7. Transfer the tissue, but not the trypsin, to another clean petri dish prepared in step 3 with 2 ml of CEM, to neutralize trypsin digestion. Mince the tissue further to 1 to 1.5 mm3 using sterile fine iris scissors, which will make the tissue ready for plating. 8. Take out the fibronectin-coated petri dishes containing 1.5 ml CEM (prepared in step 3) from the incubator and plate about 30 to 35 1- to 1.5-mm3 explants pieceby-piece on a fibronectin-coated dish with prewarmed CEM (Fig. 1A). Keep the distance between explants to 0.5 to 1 cm. Prewarming the CEM-filled fibronectin-coated dish can enhance explant adherence. The explants may not adhere properly if the CEM volume is greater than 1.5 ml.

9. Carefully replace the explant culture dishes in the incubator, trying not to dislodge explants. 10. After 24 hr, add a further 0.5 ml of CEM to the explant culture. Explants should adhere firmly to the petri dish overnight. Cells should start migrating out from the explants after 1 to 2 days. Adding more medium can prevent dehydration and improve cell growth.

11. On day 3, remove 1 ml of medium from the explant dishes and replace it with 2 ml of fresh CEM. Change all medium every 2 days after day 5. Explant-derived cells should reach 80% to 90% confluency after 7 days. Using this method, about six dishes of explants can be produced, which yield about 5–6 × 106 explant-derived cells after 7 days from one neonatal rat heart.

CARDIOSPHERE CULTURE This protocol is used to grow cardiospheres from the explant-derived cells by seeding a predetermined cell number in each well. Only loosely adherent cardiospheres are harvested to exclude adhesive non-sphere-forming cells. This method can also be used to grow secondary cardiospheres (Fig. 2C.3.1F).

BASIC PROTOCOL 2

Materials Explant-derived cells with 80% to 90% confluency (Basic Protocol 1) 2 mg/ml poly-D-lysine (Sigma, cat. no. P7280) stock solution in DPBS (store in 100-μl aliquots at –20◦ C) Dulbecco’s phosphate-buffered saline without CaCl2 and MgCl2 (DPBS, Sigma) Versene (Invitrogen) 0.05% trypsin/EDTA (Invitrogen) Complete explant medium (CEM, see recipe) Cardiosphere growth medium (CGM, see recipe) Inverted light microscope 24-well tissue culture plates (see recipe) 50-ml conical centrifuge tube Benchtop centrifuge: e.g., ALC PK121R multispeed centrifuge Hemacytometer (also see UNIT 1C.3) Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3) 1. Check the confluency of explant-derived cells under the inverted microscope. Explant confluency is preferable but not essential; it is possible to obtain sufficient cells from a nonconfluent explant that houses appreciable phase bright cells within the fibroblast outgrowth.

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A

B

C

D

E

F

Figure 2C.3.1 Cardiac stem cell isolation by explant culture. (A) Heart tissue is minced into pieces and explanted on a fibronectin-coated petri dish. (B) Phase-bright cells grow from the explant on top of a layer of stromal-like cells after 7 days. (C) Cardiospheres form after 4 days on poly-D-lysine. (D) Cardiospheres are expanded in a fibronectin-coated flask to become a monolayer. (E) Passage-0 cardiosphere-derived cells become confluent after 5 to 7 days. (F) Cardiospherederived cells form second-generation cardiospheres at passage 5. Scale bar is 250 μm.

2. Prepare poly-D-lysine working solution just before use by diluting a 100-μl aliquot of 2 mg/ml poly-D-lysine stock solution in 12 ml DPBS (without calcium and magnesium). Coat eight 24-well plates with poly-D-lysine by adding 0.5 ml of the working solution to each well and leaving the plate at 37◦ C for 30 min. Remove the solution and wash once with DPBS (without calcium and magnesium) prior to use.

Isolation and Expansion of CardiosphereDerived Stem Cells

3. Wash the explant dishes twice with 2 ml DPBS and once with 1 ml Versene (prewarm solutions to room temperature), and save all supernatant from the washes in a 50-ml conical centrifuge tube. Saving the supernatant prevents loss of loosely adherent phase-bright cells (Fig. 2C.3.1B).

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4. Add 1 ml of 0.05% trypsin/EDTA (prewarmed to 37◦ C) to each explant dishes and incubate at 37◦ C for 3 min. 5. Carefully aspirate the trypsin/EDTA solution containing the explant-derived cells and add to the 50-ml centrifuge tube. 6. Wash and quench the explant dishes with 1 ml fresh CEM (prewarmed to room temperature), carefully aspirate the CEM, and transfer to the 50-ml centrifuge tube. 7. Put 1.5 ml of CEM into the explant culture dish with the explants still in position and place it back in the incubator. The explants continue to shed cells, and isolation can be repeated up to three to four times from the same explants. Some explant tissue may detach after trypsinization. Therefore, the volume of CEM should not be more than 1.5 ml, in order to allow the explants to adhere back onto the petri dish. Add another 0.5 ml of CEM the next day and continue as per Basic Protocol 1.

8. Centrifuge the cells in the centrifuge tube 3 min at 300 × g, room temperature. 9. Discard the supernatant and resuspend the cell pellet in 2 ml of fresh CGM. 10. Count the cells with the hemacytometer under the inverted light microscope. 11. Resuspend the cells in CGM to obtain a density of 1 × 105 cells per ml. Dispense 300 μl of cells into each poly-D-lysine coated well of a 24-well plate prepared in step 2. Incubate. Explant-derived cells may aggregate and form large clusters if more than 5 × 104 cells are seeded into each well. The cell seeding density of 3 × 104 cells per well should yield approximately 500 cardiospheres without producing clumps. Cardiospheres are formed over 2 to 3 days (Fig. 1C). This protocol allows approximately 500 cardiospheres to be produced in each well. From 6 × 106 explant-derived cells, approximately 9.6 × 104 cardiospheres can be produced in 11 days from the day the explants are prepared.

CULTURE AND EXPANSION OF CARDIOSPHERE-DERIVED CELLS This protocol describes the isolation of the cardiospheres and culture of cardiospherederived cells. Cardiosphere-derived cells can be maintained in culture up to passage 5, and retain the ability to re-form cardiospheres.

BASIC PROTOCOL 3

Materials 1 mg/ml fibronectin stock solution from bovine plasma (Sigma) Dulbecco’s phosphate-buffered saline without CaCl2 and MgCl2 (DPBS, Sigma) Cardiospheres in 24-well plate (Basic Protocol 2) Complete explant medium (CEM) Trypsin, 0.05% with sodium EDTA, liquid Cardiosphere growth medium (see recipe) 25-cm2 cell culture flasks 15-ml conical centrifuge tubes (e.g., BD Falcon) 1-ml (P-1000) pipet tip Hemacytometer inverted light microscope 1. Prepare fibronectin working solution from 1 mg/ml stock as described in Basic Protocol 1, step 1. Coat a 25-cm2 cell culture flask with 1.5 ml of fibronectin working solution, leaving the flask at 37◦ C for 30 min. Discard the solution and wash once with DPBS without CaCl2 and MgCl2 prior to use.

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2. Put 0.5 to 1 ml of DPBS without CaCl2 and MgCl2 into each well. Harvest and break the cardiospheres by using 1-ml (P-1000) pipet tip and place in a sterile 15-ml centrifuge tube. Fully formed cardiospheres are semi-adherent (Fig. 2C.3.1C). Only loosely adherent cardiospheres are desired; therefore, exclude those cells or clusters that strongly adhere to the plate.

3. Centrifuge the cardiospheres in the centrifuge tube 3 min at 300 × g (1300 rpm), room temperature. 4. Discard the supernatant and resuspend the cardiosphere pellet in 7 ml of CEM. Place them in the fibronectin-coated 25-cm2 cell culture flask prepared in step 1. Incubate. Cardiospheres which are collected from one 24-well plate (approximately 1.2 × 104 in total) are seeded into one flask and labeled as passage 0. Cardiospheres will adhere to the fibronectin and grow into a monolayer of cardiosphere-derived cells (Fig. 2C.3.1D).

5. Replace the medium with fresh medium every 3 days. Once the cells reach 80% to 90% confluency, passage the cells with 0.05% trypsin (as described below) for further expansion. Cardiosphere-derived cells should form a monolayer without clusters or cardiospheres in the flask after passage 1, giving 1.5 × 107 cells at passage 2 (Fig. 2C.3.1E).

6. Wash the CDCs twice with 2 ml DPBS (without Ca and Mg) and save all supernatant from the washes in a 15-ml conical centrifuge tube. 7. Add 1 ml of 0.05% trypsin/EDTA to each culture flask and incubate at 37◦ C for 3 min. 8. Aspirate the trypsin/EDTA solution containing cardiosphere-derived cells and add to the 15-ml centrifuge tube. 9. Wash and quench the flask with 2 ml fresh CEM and transfer the CEM to the 15-ml centrifuge tube. 10. Centrifuge the cells in the centrifuge tube for 3 min at 300 × g, room temperature. 11. Discard the supernatant and resuspend the cell pellet in 3 ml of fresh CGM. 12. Count the cells with the hemacytometer under the inverted light microscope. 13. Split the cells into three fibronectin-coated 25-cm2 cell culture flasks and add 6 ml CEM to each flask. 14. Repeat steps 6 to 13 for further expansion. SUPPORT PROTOCOL

Isolation and Expansion of CardiosphereDerived Stem Cells

CHARACTERIZATION OF CARDIOSPHERE-DERIVED CELLS BY IMMUNOFLUORESCENT STAINING Cardiosphere-derived cells (CDCs) comprise a heterogeneous mixture of cells. This protocol describes immunofluorescence labeling to identify the presence of cardiac stem cells (c-kit, Oct3/4, Sox2) and differentiated cells (α-sarcomeric actin for differentiating cardiac cells, α-smooth muscle actin for smooth muscle cells, and von Willebrand factor for endothelial cells) in passage-2 CDCs following successful isolation and expansion described in Basic Protocols 1 to 3. The list of tested antibodies for this application is provided in Table 2C.3.1.

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Table 2C.3.1 Primary Antibodies Used in Characterizing Rat CDCs

Antigen

Antibody

Supplier

Catalog no.

Product identifier Dilution

CD117 (c-kit)

Rabbit polyclonal

Santa Cruz

SC-5535

H300

1:50

Smooth muscle actin

Mouse monoclonal

Sigma

A2547

1A4

1:500

Alpha-sarcomeric actin

Mouse monoclonal

Sigma

A2172

5C5

1:500

Oct 3/4

Rabbit polyclonal

Santa Cruz

SC-9081

H134

1:50

Sox 2

Goat polyclonal

Santa Cruz

SC-17320

Y17

1:50

von Willebrand factor

Rabbit polyclonal

Chemicon

AB-7356

—

1:50

Materials Fibronectin (Sigma) Rat cardiosphere-derived cells (CDCs; Basic Protocol 3) Dulbecco’s phosphate-buffered saline without CaCl2 and MgCl2 (DPBS, Sigma) 4% (w/v) paraformaldehyde Washing solution: DPBS without CaCl2 and MgCl2 , containing 0.1% (v/v) Tween 20 Permeabilization solution: DPBS without CaCl2 and MgCl2 , containing 0.1% (v/v) Tween 20 and 0.1% (v/v) Triton X-100 Blocking solution: DPBS without CaCl2 and MgCl2 , containing 0.1% (v/v) Tween 20 and 0.1% (v/v) Triton X-100 Primary and secondary antibodies (see Table 2C.3.1 for suppliers and dilution factors) 1 μg/ml DAPI (Sigma) Vectashield mounting medium (Vector Laboratories) 4-well chamber slides (Sigma) Platform shaker Humidified chamber: e.g., Tupperware box with lid containing moistened paper towels 22 × 50 mm coverslips Culture CDCs on a chamber slide 1. Prepare 10 μg/ml fibronectin and coat the 4-well chamber slides using the technique described in Basic Protocol 1. Seed about 3–5 × 104 rat CDCs per 500 μl CEM into each well. 2. Gently shake the chamber slide to make sure the cells are evenly distributed in the wells. 3. Place the chamber slide in the incubator for 4 to 6 hr, then proceed to fixation. This is to allow cells to adhere to the well, ready for fixation.

Perform cell fixation, permeabilization, and blocking 4. Remove the CEM from the chamber slide and wash three times with sterile DPBS without CaCl2 and MgCl2 . 5. Fix the cells by filling each well with cold 4% paraformaldehyde (PFA) and leave on ice for 20 min. 6. Aspirate PFA and wash the cells three times with washing solution, each time for 5 min on a platform shaker at 30 rpm.

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7. Permeabilize the cells by incubating with permeabilization solution for 10 min at room temperature. Aspirate the solution and wash the cells three times with washing solution, each time for 5 min, on a platform shaker at 30 rpm. Omit step 7 when staining for surface markers, such as c-kit (CD117).

8. Incubate each well with blocking solution for an hour at room temperature. Use the serum from the species used to raise the secondary antibody to make the blocking solution. For example, use donkey serum for donkey secondary antibody.

Perform primary antibody incubation 9. Dilute the primary antibody in DPBS without CaCl2 and MgCl2 , according to the dilution factors showed in Table 2C.3.1. 10. Aspirate the blocking solution and cover the well with the diluted primary antibody solution (∼300 μl/well). 11. Incubate the cells in the chamber slide at 4◦ C, overnight in a humidified chamber.

Perform secondary antibody labeling, DAPI staining, and mounting 12. Wash gently with DPBS without CaCl2 and MgCl2 three times, each time for 5 min, on a platform shaker at 30 rpm. 13. Add the appropriate secondary antibody solution (at the dilutions indicated in Table 2C.3.1) to each well and incubate at 37◦ C for 1 hr. Use the secondary antibody from the species used to raise the primary antibody. For example, use donkey anti-rabbit secondary antibody to label rabbit primary antibody. Recommended dilution for AlexaFluor-488 is 1:1000.

14. Wash gently with DPBS without CaCl2 and MgCl2 three times, each time for 5 min on a platform shaker at 30 rpm. 15. Stain nuclei by covering well with 1 μg/ml DAPI solution for 14 min at room temperature, in a humidified chamber and protected from light. Omit this step if using the Vectashield mounting medium which contains DAPI or propidium iodide.

16. Wash gently with DPBS without CaCl2 and MgCl2 three times, each time for 5 min, on a platform shaker at 30 rpm. 17. Remove the chamber with a slide separator (provided by the manufacturer). Do exactly as instructed by the manufacturer. Inappropriate removal of the chamber may break the slide.

18. Put a drop of Vectashield mounting medium on each well/rectangle and cover the sample with a 22 × 50–mm cover glass. Visualize the staining with a fluorescent/confocal microscope.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX. All reagents and solutions below are prepared under sterile conditions.

Cardiosphere-growth medium (CGM) Isolation and Expansion of CardiosphereDerived Stem Cells

65% (v/v) Dulbecco’s modified Eagle’s medium (DMEM)/F12 (Invitrogen)/35% (v/v) Iscove’s modified Dulbecco’s medium (IMDM; Invitrogen), supplemented with: continued

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7% (v/v) fetal bovine serum (Biosera) 1× penicillin-streptomycin-glutamine (added from 100× solution which contains 10,000 U/ml penicillin, 10,000 μg/ml streptomycin, and 29.2 mg/ml Lglutamine/ml in 0.85% saline; Invitrogen) 0.1 mM 2-mercaptoethanol (Sigma) 2% (v/v) B27 supplement (Invitrogen) 25 ng/ml cardiotrophin (see recipe for stock solution) 20 ng/ml bFGF (see recipe for stock solution) 10 ng/ml EGF (see recipe for stock solution) 5 U thrombin (see recipe for stock solution) To make CGM, add one aliquot of each growth factor/cytokine to 20 ml of 35% IMDM and 65% DMEM/F12 with 7% FBS. Store up to 2 weeks at 4◦ C

Complete explant medium (CEM) Iscove’s modified Dulbecco’s medium (IMDM) supplemented with: 20% (v/v) fetal bovine serum (FBS; Biosera, http://www.biosera.com/) 1× penicillin-streptomycin-glutamine (added from 100× solution which contains 10,000 U/ml penicillin, 10,000 μg/ml streptomycin, and 29.2 mg/ml Lglutamine/ml in 0.85% saline; Invitrogen) 0.1 mM 2-mercaptoethanol (Sigma) Store for up to 8 weeks at 4◦ C Human recombinant cardiotrophin-1 stock Dissolve 10 μg of cardiotrophin (Peprotech) in 100 μl of 20 mM Tris·Cl, pH 8.0. Store the stock solution in 5-μl aliquots at −20◦ C.

Human recombinant epidermal growth factor (EGF) stock Dissolve 100 μg of EGF (Peproptech) in 200 μl sterile 0.1% (w/v) BSA in doubledistilled water. Store EGF concentrates in 20-μl aliquots at −20◦ C. Dilute each concentrate to 500 μl and store it in 10-μl aliquots at −20◦ C.

Thrombin stock Dissolve 100 U thrombin from human plasma (Sigma) in 1 ml DPBS without CaCl2 and MgCl2 and store in 50-μl aliquots at −20◦ C.

Recombinant human basic fibroblast growth factor (bFGF) Dissolve 25 μg of bFGF (Promega) in 250 μl of serum-free DMEM/F12 medium (Invitrogen). Store the stock solution in 4-μl aliquots at −20◦ C.

COMMENTARY Background Information Even with the best available therapy, myocardial infarction can progress to heart failure and death (Cowie et al., 2000; also see Internet Resources). Current pharmacological therapies are able to alleviate short-term cardiac function, but fail to address the fundamental problems: substantial reduction in myocyte number and high levels of noncontractile scar tissue. To solve this, attention has shifted to stem cell therapy, with the aim of reconstituting the scar with viable, functional

cardiomyocytes and vasculature. Several cell types have been proposed for cardiac regeneration, including skeletal myoblasts (Menasche, 2007), bone-marrow derived cells (Orlic, 2003), and embryonic stem cells (Murry and Keller, 2008). However, recently identified resident cardiac stem cells, believed to be responsible for regulating cardiac homeostasis and myocyte turnover (Torella et al., 2006), may be the most suitable choice for autologous cell therapy for the heart (Barile et al., 2007).

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Isolation and Expansion of CardiosphereDerived Stem Cells

Endogenous cardiac stem cells are selfrenewing, clonogenic, and multipotent—able to undergo differentiation into cardiomyocytes, smooth muscle, and endothelial cells (Beltrami et al., 2003; Oh et al., 2003; Tateishi et al., 2007) or transdifferentiation into adipocytes and osteocytes (Matsuura et al., 2004). Various markers have been used to identify putative cardiac stem cells such as c-kit, Sca-1, or Isl-1, and methods of isolation have been reported using magnetic (Tang et al., 2009) or fluorescent cell sorting (Itzhaki-Alfia et al., 2009), or both (Beltrami et al., 2003; Oh et al., 2003), or single-cell clonogenic amplification (Tateishi et al., 2007). However, as the number of stem cells present in the heart is small, isolation based on specific cell-surface markers yields relatively few cells and requires substantial in vitro expansion to generate sufficient cell numbers for therapy. This is important, as the efficacy of cell therapy appears to directly relate to the number of cells grafted and inversely correlates with time of cell delivery after reperfusion therapy post infarction (Schachinger et al., 2006; Lipinski et al., 2007). Messina et al. (2004) developed a technique of tissue explant culture and cell expansion via the formation of cardiospheres using defined growth factors. The resulting cells can be expanded in a monolayer, and express the stem cell marker, c-kit (CD117), the mesenchymal marker Thy-1 (CD90), and endoglin (CD105), one of the components of the transforming growth factor-β receptor complex important in angiogenesis. This method is capable of generating therapeutically relevant numbers of cells in a shorter time period than cell surface marker–based methods. Smith et al. (2007) adapted the technique, expanding cardiospheres to CDCs as a monolayer, and generated 2 × 106 human CDCs over 45 days, with the cells expressing c-kit, CD90, CD105, CD34, and CD31. We have modified Smith’s technique to optimize cell density (3 × 104 cells/well) and serum concentration (7%) to grow cardiospheres, thereby yielding 1.5 × 107 rat CDCs in 16 days, which is a sufficient number for cell therapy application. We found that rat neonatal CDCs cultured using this protocol expressed the pluripotent markers Oct 3/4 and SOX2, and cardiac lineage specific markers α-sarcomeric actin and von Willebrand factor (Figure 2C.3.2). The explant technique also enables isolation of high numbers of stem cells from small human biopsies.

Critical Parameters and Troubleshooting CDC culture consists of three steps (explantation, cardiosphere culture and expansion of CDCs), each step involving different media and methods. Explant culture: Explants should be kept to around 1 to 1.5 mm3 in size. Larger explants can easily detach from the dish, and no cells will grow out from an unattached explant. To facilitate explant adherence, the volume of the culture medium in the explant dish must be kept to a minimum, but enough to cover the explants and a thin layer of the petri dish on the first day. Prewarming the medium in the fibronectin-coated petri dish to 37◦ C can also improve explant adherence. Cardiosphere culture: To grow cardiospheres, the cell density in each well is critical. Cardiospheres grow very slowly if seeded at a low density, whereas high cell seeding density will cause the cells to clump and form large aggregates. CDC culture: CDCs can be passaged every 5 to 6 days. CDCs may form secondary and tertiary cardiospheres following Basic Protocol 2.

Anticipated Results This protocol allows large-scale expansion of cardiosphere-derived cells from heart tissue. One neonatal rat heart can produce six dishes of explants, which should yield 5–6 × 106 explant-derived cells that can be seeded into at least eight plates of a 24-well plate to grow cardiospheres. Approximately 500 cardiospheres are produced in each well (resulting in a total of 9.6 × 104 cardiospheres), and can be expanded as monolayers in eight 25-cm2 flasks. This gives 1.5 × 107 CDCs at the end of passage 0. CDCs cultured to passage 2 have been shown to improve function when injected into the infarcted heart (Smith et al., 2007).

Time Considerations Explant-derived cells may be harvested from neonatal heart explants after 7 days. Older rat hearts and human heart pieces may need longer for cells to migrate from the explants (up to 14 days). The explant-derived cells are harvested and produce 500 cardiospheres over 3 to 4 days. Cardiospheres are expanded as a monolayer on fibronectin coated flasks and reach 90% confluency after 6 to

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A

c-Kit Propidium lodide

B

SOX2 DAPI

50 m

C

Oct-4 DAPI

D

50

m

-sarcomeric actin DAPI

50

m

F

E

Smooth muscle actin DAPI

50 m

50 m

vWF DAPI

50 m

Figure 2C.3.2 Characterization of cardiosphere-derived cells (CDCs). The CDCs comprised a mixed cell population consisting of c-kit-expressing (green) cardiac stem/progenitor cells (A), undifferentiated, more primitive progenitors expressing SOX2 (red) and Oct-4 (green) (B and C), cardiomyocyte-specific committed cells (green, α-sarcomeric actin; D), smooth muscle cells (green, smooth muscle actin; E), and endothelial cells (green, von Willebrand factor (vWF); F). Nuclei are stained with propidium iodide in red (A) and by DAPI in blue (B-F). Scale bar is 50 μm.

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7 days. Thus, this protocol allows 1.5 × 107 CDCs to be produced in 16 days.

Acknowledgements Our work is funded by the British Heart Foundation and the Malaysian Ministry of Higher Education.

Literature Cited Barile, L., Messina, E., Giacomello, A., and Marban, E. 2007. Endogenous cardiac stem cells. Prog. Cardiovasc. Dis. 50:31-48. Beltrami, A.P., Barlucchi, L., Torella, D., Baker, M., Limana, F., Chimenti, S., Kasahara, H., Rota, M., Musso, E., Urbanek, K., Leri, A., Kajstura, J., Nadal-Ginard, B., and Anversa, P. 2003. Adult cardiac stem cells are multipotent and support myocardial regeneration. Cell 114:763776. Cowie, M.R., Wood, D.A., Coats, A.J.S., Thompson, S.G., Suresh, V., Poole-Wilson, P.A., and Sutton, G.C. 2000. Survival of patients with a new diagnosis of heart failure: A population based study. Heart 83:505-510. Davis, D.R., Ruckdeschel Smith, R., and Marban, E. 2010. Human cardiospheres are a source of stem cells with cardiomyogenic potential. Stem Cells 28:903-904. Itzhaki-Alfia, A., Leor, J., Raanani, E., Sternik, L., Spiegelstein, D., Netser, S., Holbova, R., Pevsner-Fischer, M., Lavee, J., and Barbash, I.M. 2009. Patient characteristics and cell source determine the number of isolated human cardiac progenitor cells. Circulation 120:25592566. Lipinski, M.J., Biondi-Zoccai, G.G., Abbate, A., Khianey, R., Sheiban, I., Bartunek, J., Vanderheyden, M., Kim, H.S., Kang, H.J., Strauer, B.E., and Vetrovec, G.W. 2007. Impact of intracoronary cell therapy on left ventricular function in the setting of acute myocardial infarction: A collaborative systematic review and metaanalysis of controlled clinical trials. J. Am. Coll. Cardiol. 50:1761-1767. Matsuura, K., Nagai, T., Nishigaki, N., Oyama, T., Nishi, J., Wada, H., Sano, M., Toko, H., Akazawa, H., Sato, T., Nakaya, H., Kasanuki, H., and Komuro, I. 2004. Adult cardiac Sca-1-positive cells differentiate into beating cardiomyocytes. J. Biol. Chem. 279:1138411391. Menasche, P. 2007. Skeletal myoblasts as a therapeutic agent. Progr. Cardiovasc. Dis. 50:7-17. Messina, E., De Angelis, L., Frati, G., Morrone, S., Chimenti, S., Fiordaliso, F., Salio, M., Battaglia,

M., Latronico, M.V., Coletta, M., Vivarelli, E., Frati, L., Cossu, G., and Giacomello, A. 2004. Isolation and expansion of adult cardiac stem cells from human and murine heart. Circulation Res. 95:911-921. Murry, C.E. and Keller, G. 2008. Differentiation of embryonic stem cells to clinically relevant populations: Lessons from embryonic development. Cell 132:661-680. Oh, H., Bradfute, S.B., Gallardo, T.D., Nakamura, T., Gaussin, V., Mishina, Y., Pocius, J., Michael, L.H., Behringer, R.R., Garry, D.J., Entman, M.L., and Schneider, M.D. 2003. Cardiac progenitor cells from adult myocardium: homing, differentiation, and fusion after infarction. Proc. Natl. Acad. Sci. U.S.A. 100:12313-12318. Orlic, D. 2003. Adult bone marrow stem cells regenerate myocardium in ischemic heart disease. Ann. New York Acad. Sci. :152-157. Schachinger, V., Erbs, S., Elsasser, A., Haberbosch, W., Hambrecht, R., Holschermann, H., Yu, J., Corti, R., Mathey, D.G., Hamm, C.W., S¨uselbeck, T., Assmus, B., Tonn, T. Dimmeler, S., and Zeiher, A.M. 2006. Intracoronary bone marrow-derived progenitor cells in acute myocardial infarction. N. Engl. J. Med. 355:12101221. Smith, R.R., Barile, L., Cho, H.C., Leppo, M.K., Hare, J.M., Messina, E., Giacomello, A., Abraham, M.R., and Marban, E. 2007. Regenerative potential of cardiosphere-derived cells expanded from percutaneous endomyocardial biopsy specimens. Circulation 115:896-908. Tang, Y.L., Zhu, W., Cheng, M., Chen, L., Zhang, J., Sun, T., Kishore, R., Phillips, M.I., Losordo, D.W., and Qin, G. 2009. Hypoxic preconditioning enhances the benefit of cardiac progenitor cell therapy for treatment of myocardial infarction by inducing CXCR4 expression. Circulation Res. 104:1209-1216. Tateishi, K., Ashihara, E., Honsho, S., Takehara, N., Nomura, T., Takahashi, T., Ueyama, T., Yamagishi, M., Yaku, H., Matsubara, H., and Oh, H. 2007. Human cardiac stem cells exhibit mesenchymal features and are maintained through Akt/GSK-3beta signaling. Biochem. Biophys. Res. Commun. 352:635-641. Torella, D., Ellison, G.M., Mendez-Ferrer, S., Ibanez, B., and Nadal-Ginard, B. 2006. Resident human cardiac stem cells: Role in cardiac cellular homeostasis and potential for myocardial regeneration. Nat. Clin. Practice 3:S8-S13.

Internet Resources http://www.heartstats.org/datapage.asp?id=752 Online version of Cowie et al. (2000); also see Literature Cited.

Isolation and Expansion of CardiosphereDerived Stem Cells

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Current Protocols in Stem Cell Biology

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

UNIT 2D.1

Eric C. Larsen,1 Yoichi Kondo,1 Cale D. Fahrenholtz,1 and Ian D. Duncan1 1

Department of Medical Sciences, University of Wisconsin-Madison, Madison, Wisconsin

ABSTRACT The oligodendrocyte progenitor cell (OPC) is one of the most studied progenitor cells of the body. It has been extensively researched in tissue culture and more recently in vivo using a wide range of markers that recognize transcription factors and cell surface markers and identify its earliest development from neural stem cells onward. Isolation of OPCs in large numbers and in purified preparations has been sought after as a source of cells for the repair of human myelin disorders. It has been proposed that such cells could be used as an exogenous source of cells for the treatment of lesions in multiple sclerosis and the less common genetic myelin disorders such as Pelizaeus-Merzbacher disease. Prior to clinical trials, such approaches can be tested in animal models. Here, we describe the isolation of rat OPCs in culture conditions that provide large numbers of purified C 2008 by John populations of cells. Curr. Protoc. Stem Cell Biol. 6:2D.1.1-2D.1.13.  Wiley & Sons, Inc. Keywords: oligodendrocytes r oligodendrocyte progenitor cells r oligospheres r neurospheres r differentiation r transplantation

INTRODUCTION We describe in this unit a protocol for the isolation and culturing of oligodendrocyte progenitor cells (OPCs) from the brains of rat neonates. Once OPCs are removed from neural tissue and placed in culture medium preconditioned by B104 neuroblastoma cells, they will form free-floating, spherical clusters of cells called oligospheres (Fig. 2D.1.1A; Basic Protocol), a term first coined by Avellana-Adalid et al. (1996). These oligospheres continue to proliferate in culture for months after their isolation from the rat neonatal brain and provide a readily available source of OPCs. In addition, we have included an Alternate Protocol describing how to generate OPCs from neural precursor cells. Furthermore, three support protocols have been included. The first describes the production of B104conditioned medium (Support Protocol 1), which is an essential component of oligosphere medium. The remaining protocols detail the in vitro differentiation (Support Protocol 2) and transplantation (Support Protocol 3) of cultured rat OPCs, respectively. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. NOTE: All surgical equipment should be sterilized prior to usage.

Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.1.1-2D.1.13 Published online August 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d01s6 C 2008 John Wiley & Sons, Inc. Copyright 

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Figure 2D.1.1 Phase contrast images of oligodendrocyte progenitor cells (OPCs) in culture. (A) Once cultured from rat neonatal brain, OPCs will typically form floating clusters of cells called oligospheres. (B) On some occasions, oligospheres will attach to the surface of the culture flask; when this happens, individual cells will be observed migrating away from the oligosphere. (C) Isolated OPCs will typically adopt a bipolar morphology with little secondary branching. (D) As OPCs differentiate, more processes will emerge from the cell body, and more secondary branching will be observed coming from the processes.

BASIC PROTOCOL

GENERATION OF OPC CULTURES FROM RAT NEONATES The protocol for the generation of rat OPC cultures has been divided into two parts. The first part describes the isolation of OPC cultures from the brains of rat neonates. The second part details the maintenance of the cultured OPCs and subsequent formation of oligospheres.

Materials Rat neonates (post-natal day 0 to 6) Isoflurane (Abbot, cat. no. 5260-04-05) 70% ethanol Hank’s Balanced Salt Solution (HBSS), without Ca2+ and Mg2+ (Invitrogen, cat. no. 14175) Oligosphere medium (see recipe) Sphere freezing medium: 80% DMEM/10% FBS/10% DMSO Liquid nitrogen

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

Small, airtight container Towel Class II biological hazard flow hood or laminar-flow hood Surgical scissors, sterile (Fine Science Tools, cat. no. 14090-11) Forceps, sterile (no. 5 and no. 55; Fine Science Tools, cat. nos. 11252-20 and 11255-20) Rat tooth forceps, sterile (Fine Science Tools, cat. no. 11022-14)

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60-mm and 100-mm tissue culture dishes, sterile (Nunc, cat. nos. 150288 and 172958) Dissecting microscope Curved spring scissors, sterile (Fine Science Tools, cat. no. 15023-10) 15-ml tube, sterile (Fisher, cat. no. 05-539-5) Fire-polished Pasteur pipets (Fisher, cat. no. 13-678-20D) 25-cm2 flasks, sterile (Nunc, cat. no. 136196) Humidified incubator set at 37◦ C and 5% CO2 Cryovial (Nunc, cat. no. 375418) Freezing container (Nunc, cat. no. 5100-0001) −80◦ C freezer 37◦ C water bath Remove neonate brain 1. Place one pup in a small, airtight container containing an isoflurane-soaked towel. Euthanize by isoflurane inhalation. 2. Transfer the pup to the laminar-flow hood, spray the pup with 70% ethanol, and decapitate it with a sharp pair of sterile surgical scissors. 3. Expose the skull by cutting the skin from the neck to the nose with surgical scissors. 4. With a new pair of sterile surgical scissors and sterile forceps, remove the crown of the skull to expose the brain. 5. Remove the brain by gently slipping rat tooth forceps underneath the brain and teasing the brain out to minimize tissue disruption. 6. Place the brain in a 60-mm dish containing a small amount of HBSS.

Dissect the brain 7. Under a dissecting microscope, remove and discard the cerebellum, olfactory bulbs, and meninges. 8. Using a pair of curved spring scissors, cut the cerebral cortex from front to back. 9. Expose the striatum by gently peeling back the cortex with forceps. 10. Gently ease the striatum out from each hemisphere and place in a 100-mm dish containing ∼10 ml HBSS on ice. 11. Repeat steps 1 to 10 for each remaining pup, placing each dissected striatum into the same 100-mm dish. Typically, we only pool striata from pups in the same litter.

Isolate cells 12. Transfer the contents of the dish into a 15-ml tube. Centrifuge 2 min at 225 × g, room temperature. Aspirate the supernatant. 13. Resuspend the tissue in ∼2 ml oligosphere medium. 14. Using prewetted fire-polished Pasteur pipets with a succession of larger-bore to smaller-bore openings, thoroughly triturate the tissue to pieces smaller than 1 mm3 . If the procedure described in step 14 is insufficient in breaking up the tissue into small enough pieces, several approaches may be utilized. Tissue may be pretreated prior to trituration by resuspending the tissue from step 12 in ∼1 ml 0.25% trypsin/EDTA (Invitrogen, cat. no. 25200), followed by incubation at 37◦ C for 15 min and addition of 1 ml heatinactivated normal horse serum (Invitrogen, cat. no. 26050-070). Alternatively, tissue may be minced by hand with a sterile razor blade or surgical scalpel into smaller tissues following step 11 and then processed accordingly. Similarly, the cell suspension may also

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be filtered through a 100-μm cell strainer (BD Falcon, cat. no. 352360), although doing so will result in some tissue loss.

15. Calculate the total volume of the cell suspension and distribute it equally among sterile 25-cm2 flasks containing 4.5 ml oligosphere medium that has been equilibrated for at least 15 min in a humidified, 37◦ C, 5% CO2 incubator. Typically, we use one 25-cm2 flask per pup.

Culture and maintain oligospheres 16. Following isolation from the rat neonate brain, incubate the cultured cells in an incubator set at 37◦ C and 5% CO2 . 17. Every 2 to 3 days, replace approximately half of the medium with fresh oligosphere medium. a. Remove the flask from the incubator, gently tap the flask to remove attached spheres, and tilt the flask to allow spheres to settle to the bottom. b. Remove half of the medium in the flask, taking care not to remove spheres along with the medium. c. Replace with an equivalent volume of fresh, prewarmed oligosphere medium and return to the incubator. 18. Continue culture for ∼4 weeks, by which time the cultures will contain primarily oligospheres. While oligosphere cultures do not need to be split per se, oligospheres do need to be broken up into smaller aggregates when they reach a size >100 μm. Depending on the growth of the culture, this may need to be performed every 1 to 2 weeks. To break up oligospheres, triturate using a series of fire-polished pipets, ranging from larger-bore to smaller-bore openings. Mild trypsinization can be used prior to trituration if oligospheres are not easily broken up.

19. Continue to culture oligospheres, which will remain viable in culture for months. Alternatively, freeze oligospheres and then thaw for use at a later date. a. To freeze oligospheres, transfer spheres to a 15-ml tube and centrifuge for 7 min at 225 × g, room temperature. Gently resuspend the pellet in sphere freezing medium and transfer to a cryovial. Place cryovials in a freezing container overnight in a −80◦ C freezer. The next day, transfer vials to liquid nitrogen. b. To thaw frozen oligospheres, thaw vials rapidly in a 37◦ C water bath with gentle swirling. Transfer the contents of the vials to a 15-ml tube and adjust volume to 10 ml with oligosphere medium. After gentle mixing, centrifuge 4 min at 200 × g, room temperature. Gently resuspend the pellet in 5 ml medium and transfer to a flask. Incubate at 37◦ C/5% CO2 and allow several days for recovery. While the majority of oligospheres will remain floating in the culture medium, in some cases oligospheres will attach to the bottom of the culture flask. When they do so, individual cells will migrate away from the attached oligosphere (Fig. 2D.1.1B). ALTERNATE PROTOCOL

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

GENERATION OF OLIGOSPHERES FROM NEURAL PRECURSOR CELLS The Basic Protocol outlined in this unit describes the most direct method for generating OPCs from the brains of rat neonates. In some situations, however, it is desirable to initially generate cultured neural precursor cells from rat neonatal brains. Neural precursor cells, which typically aggregate into neurospheres when cultured, are capable of differentiating into neurons, given the appropriate conditions. Alternatively, neurospheres can be converted into oligospheres. Neurospheres have an advantage in that they can be converted either into neuronal or glial cell types, whereas oligospheres are restricted

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to a glial cell fate. The procedure for this neurosphere-to-oligosphere conversion was first described in Zhang et al. (1998a). The protocol involves the gradual replacement of neurosphere medium with oligosphere medium, which will lead to the conversion of neurospheres into oligospheres.

Additional Materials (also see Basic Protocol) Neurosphere medium (see recipe) 1. Isolate neural precursor cells using Basic Protocol steps 1 to 15, with the exception of using neurosphere medium in place of oligosphere medium. After ∼4 weeks in neurosphere medium, the culture preparation will contain primarily neurospheres.

2. Remove the flask containing neurospheres from the incubator, tap the flask to dislodge any attached neurospheres, and tilt the flask to allow neurospheres to settle to the bottom. 3. Remove approximately half the medium in the flask, taking care not to remove spheres along with the neurosphere medium. 4. Replace with an equivalent volume of prewarmed oligosphere medium and return to the incubator. 5. Every 2 to 3 days, repeat steps 2 and 3, using fresh oligosphere medium each time. After ∼4 weeks of medium exchange, the neurospheres will be converted to oligospheres.

PRODUCTION OF B104-CONDITIONED MEDIUM An essential component of the medium used to culture oligospheres is medium that has been preconditioned by exposure to B104 neuroblastoma cells (first described by Avellana-Adalid et al. in 1996). The factors present in B104-conditioned medium that enhance the growth of oligospheres remain unknown, although growth factors such as transforming growth factor P (TGF-P) and platelet-derived growth factor (PDGF) are among the potential candidates (Asakura et al., 1997). This support protocol describes the procedure for producing B104-conditioned medium to be used in oligosphere medium.

SUPPORT PROTOCOL 1

Materials B104 neuroblastoma cells (generously provided by Dr. M. Dubois-Dalcq) B104 feeding medium (see recipe) Trypsin/EDTA (Invitrogen, cat. no. 25200) Trypan blue (Invitrogen, cat. no. 15250-061) B104 collection medium (see recipe) 37◦ C water bath Sterile 15-ml and 50-ml tubes (Fisher, cat. nos. 05-539-5 and 05-539-8, respectively) Sterile 75-cm2 (TPP, cat. no. 90076) and 175-cm2 (Corning, cat. no. 431080) culture flasks Hemacytometer (Hausser Scientific, cat. no. 1490) 0.22-μm filter (Millipore, cat. no. SCGPU05RE) Additional reagents and equipment for performing a viable cell count using a hemacytometer and trypan blue (UNIT 1C.3) Start B104 cultures 1. On day 1, thaw B104 cells (1 × 106 cells) rapidly in a 37◦ C water bath. Transfer contents to a sterile 15-ml tube.

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2. Using B104 feeding medium, adjust volume to 10 ml and mix gently. 3. Centrifuge 8 min at 300 × g, room temperature. 4. Aspirate the supernatant and resuspend the pellet in 2 ml B104 feeding medium. 5. Transfer the cell suspension to a sterile 75-cm2 flask containing 10 ml feeding medium. 6. The next day (day 2), exchange medium with fresh B104 feeding medium.

Passage cells 7. The following day (day 3), when the flask is confluent with B104 cells, aspirate medium and add 3 ml trypsin/EDTA. 8. Once the B104 cells have become detached from the flask, add 7 ml B104 feeding medium and mix gently. 9. Transfer the cell suspension to a sterile 15-ml tube. Count viable cells using a hemacytometer and trypan blue (UNIT 1C.3). 10. Plate 45,000 cells in sterile 175-cm2 flasks containing 25 ml B104 feeding medium. 11. After 4 days (day 7), aspirate the feeding medium from the 175-cm2 flasks. Replace with 25 ml B104 collection medium.

Collect medium 12. After 3 days in collection medium (day 10), transfer all medium to sterile 50-ml tubes. Discard B104 cells. 13. Centrifuge 10 min at 300 × g, room temperature to remove cell debris. 14. Filter-sterilize medium through a 0.22-μm filter. Divide into aliquots and store at −20◦ C until use. This medium is now B104-conditioned medium. SUPPORT PROTOCOL 2

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

IN VITRO DIFFERENTIATION OF CULTURED OPCS Once cultured, oligospheres can be subsequently differentiated in vitro. It should be noted, however, that oligosphere differentiation rarely results in a completely homogeneous population of oligodendrocytes. Rather, a mixed population of oligodendrocytes and astrocytes will typically be generated by in vitro oligosphere differentiation. Even with this caveat, in vitro differentiation of oligospheres provides an incredibly useful tool for studying oligodendrocyte function that is not possible in vivo. Oligodendrocyte differentiation from oligodendrocyte progenitor cells is marked by significant changes in cell morphology and antigenicity. Isolated OPCs initially have a bipolar morphology, with little or no secondary branching (Fig. 2D.1.1C). As OPCs differentiate and mature, more processes emerge from the cell body, and more significant branching from these processes is observed (Fig. 2D.1.1D). Likewise, changes in cell antigenicity can be used to follow the differentiation process (Fig. 2D.1.2). Isolated OPCs are characterized as being positive for A2B5 and the α receptor for platelet-derived growth factor (PDGFαR). As OPCs differentiate into pre-oligodendrocytes (pre-oligo), O4 antigenicity will be observed in addition to A2B5 and PDGFαR antigenicity. As cells continue to differentiate into immature, premyelinating oligodendrocytes (immature oligo), A2B5 and PDGFαR antigenicity is lost, and cells will become positive for galactosylceramidase (GalC). Finally, mature, myelinating oligodendrocytes (mature oligo) will stain for the myelin proteins myelin basic protein (MBP) and proteolipid protein (PLP). This support protocol details the process for differentiating cultured OPCs.

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Antigen expression

OPC

immature oligo

pre-oligo

O4

GalC

mature oligo

MBP, PLP

Time in culture Figure 2D.1.2 In addition to morphological changes, changes in cell antigenicity can be observed as cultured OPCs differentiate into mature oligodendrocytes. OPCs will be positive for A2B5 and PDGFαR. OPCs will first differentiate into pre-oligodendrocytes (pre-oligo) and will be positive for O4 in addition to A2B5/PDGFαR. Continued differentiation into immature, nonmyelinating oligodendrocytes results in a loss of antigenicity for A2B5/PDGFαR and a gain of antigenicity for galactosylceramidase (GalC). Complete maturation into myelinating oligodendrocytes results in antigenicity for the myelin proteins myelin basic protein (MBP) and proteolipid protein (PLP).

Materials Oligospheres (from the Basic Protocol or Alternate Protocol) Hank’s Balanced Salt Solution (HBSS), without Ca2+ and Mg2+ 2 mg/ml bovine serum albumin (BSA; Sigma, cat. no. A-7906) in HBSS (Invitrogen, cat. no. 14175) Oligosphere differentiation medium (see recipe) Trypan blue (Invitrogen, cat. no. 15250-061) 15-ml tubes, sterile Fire-polished Pasteur pipets Hemacytometer (Hausser Scientific, cat. cat. no. 1490) Additional reagents and equipment for performing a viable cell count using a hemacytometer and trypan blue (UNIT 1C.3) Collect oligospheres 1. Collect oligospheres and transfer to a 15-ml tube. 2. Bring to a volume of 10 ml with HBSS and mix gently using a pipet. 3. Centrifuge 5 min at 225 × g, room temperature. 4. Resuspend spheres in 2 ml HBSS. 5. Triturate the oligospheres using prewetted fire-polished Pasteur pipets with a succession of larger-bore to smaller-bore openings. 6. Allow the cell suspension to settle 1 to 2 min to allow unbroken spheres to settle. Gently transfer the single-cell suspension to a new 15-ml tube. Be careful not to disturb any unbroken spheres. Trypsinization can aid in the disruption of oligospheres. Resuspend spheres in 2 ml trypsin after step 3. Incubate in a 37◦ C water bath for 5 min, then triturate as described in step 5. Add 200 μl fetal bovine serum and mix, allow the cell suspension to settle, and transfer the single-cell suspension to a new 15-ml tube. Adjust volume to 5 ml with HBSS, then centrifuge 7 min at 225 × g, room temperature. Aspirate the supernatant, resuspend pellet in 3 ml HBSS, and proceed to step 8. Somatic Stem Cells

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Clean up the cells 7. Bring the cell suspension to a final volume of 3 ml with HBSS. Mix gently using a pipet. 8. Gently layer the cell suspension on top of a solution of 2 mg/ml BSA in HBSS. 9. Centrifuge 10 min at 130 × g, room temperature. This centrifugation will remove much of the cellular debris and dead cells that result from the dissociation of spheres into single cells.

10. Aspirate the supernatant. Resuspend the pellet, which will contain single viable cells, in 1 to 5 ml oligosphere differentiation medium. Use the smallest volume necessary to obtain a reliable cell count while keeping the concentration high enough for plating needs.

11. Count viable cells in a 10-μl aliquot using a hemacytometer and trypan blue (UNIT 1C.3). 12. Plate cells at a density of ∼150 cells/mm2 of surface area. If plating cells onto coverslips for immunostaining, precoat coverslips (Bellco, cat. no. 1943-10012) with poly-L-ornithine (Sigma, cat. no. P-3655). Plate cells (in a volume of no more than 50 μl) on coated coverslips (in a 12-well plate) and incubate 30 to 45 min in a humidified incubator set at 37◦ C and 5% CO2 to allow cells to attach. Then, carefully add 0.5 to 1.0 ml oligosphere differentiation medium. Alternatively, cells can be plated directly onto chamber slides (Nunc, cat. no.154526). SUPPORT PROTOCOL 3

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

TRANSPLANTATION OF CULTURED OPCS In this protocol we will describe how to prepare rat oligospheres for transplantation, as well as the transplantation protocol itself. Oligospheres must first be dissociated into single-cell suspensions prior to transplantation; this dissociation protocol is similar to that described in Support Protocol 1. OPC suspensions can then be transplanted into the white matter of the rat CNS. When injected into rat models of myelin disease, OPCs are typically injected into the brain at post-natal day 0 to 1 and into the dorsal column of the spinal cord at post-natal day 5 to 7 (Tontsch et al., 1994; Utzschneider et al., 1994; Zhang et al., 2003). When transplanted into rat models of multiple sclerosis, OPCs are typically injected into the dorsal column of the thoracolumbar spinal cord of adult rats at various time points of the EAE disease course.

Additional Materials (also see Support Protocol 2) Crushed ice 0.5-ml microcentrifuge tube Gauze Pulled glass micropipets (see recipe) Programmable syringe pump (Kent Scientific, cat. no. GENIE) Heating pad Isoflurane anesthesia system (including vaporizer and O2 cylinders) Stereotaxic frame (Stoelting, cat. no. 51600) Spring scissors (Fine Scientific Tools, cat. no. 15023-10) Bone-cutting spring scissors (Fine Scientific Tools, cat. no. 16144-13) High-speed microdrill (Fine Scientific Tools, cat. no. 18000-17) 0.5-mm diameter steel burr (Fine Scientific Tools, cat. no. 19007-05) Surgical spade 31-G insulin syringe (BD, cat. no. 328468; bend the needle tip with a needle holder such that the needle has an angle of ∼90-120◦ ) Micromanipulator

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Dissociate oligospheres 1. Perform steps 1 to 9 as described in Support Protocol 2. 2. Following aspiration of the supernatant produced by centrifugation (Support Protocol 2, step 9), resuspend the pellet in 1 ml HBSS. 3. Count viable cells in a 10-μl aliquot using a hemacytometer and trypan blue (UNIT 1C.3). 4. Centrifuge the cells 8 min at 225 × g, room temperature. Aspirate the supernatant. 5. Resuspend the cell pellet in an appropriate volume of HBSS to yield a cell concentration of 50,000 to 100,000 viable cells/μl. Transfer the resuspended cells to a sterile 0.5-ml microcentrifuge tube and store on ice until transplantation.

Transplant into rat neonatal brain 6a. Wrap a rat neonate (P0 to P1) in gauze and cover in crushed ice. Once the pup displays pale skin, no movement, and no respiration, it has been properly cryoanesthetized. This process will take ∼2 min, although this time may need to be optimized for each laboratory.

7a. Load a pulled glass micropipet with 2 μl of suspended OPCs (50,000 to 100,000 viable cells/μl). 8a. Insert the glass micropipet into the brain such that the OPCs will be injected into the third ventricle. Rat OPCs injected at this site are capable of migrating throughout the CNS parenchyma (Learish et al., 1999).

9a. Using a programmable syringe pump, inject the cell suspension at a rate of 2.00 μl/min. 10a. Leave the micropipet in the brain for 30 sec after injection (to prevent the backflow of cells), then slowly withdraw the micropipet. 11a. Place the pup on a heating pad until it regains consciousness, then return it to its mother. Transplant into rat spinal cord 6b. Anesthetize rat with isoflurane gas and place it onto a stereotaxic frame. 7b. Make a dorsal midline skin incision. Clear any muscle or other tissue in order to have access to the spinal cord. 8b. Remove the spinous process at the thoracolumbar level. For a rat pup (approximately P7), remove the spinous process by cutting the lamina of the vertebrae with a small pair of spring scissors. For a young/adult rat, cut the lamina with a pair of bone-cutting spring scissors. To aid in this process, use a microdrill with a 0.5-mm burr.

9b. Expose the dorsal column of the spinal cord and clear the surface with a surgical spade. Take care not to damage the dorsal spinal artery, as bleeding from the artery will require clearing of the surgical field.

10b. With a bent 31-G needle, cut a short length of the dura at the injection site. 11b. Load a pulled glass micropipet with an appropriate volume of suspended OPCs (50,000 to 100,000 viable cells/μl). For neonatal pups, inject 1 μl of cell suspension; inject 2 μl of cell suspension into the spinal cord of young or adult rats.

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12b. Using a micromanipulator, insert the micropipet containing the OPC cell suspension to the depth of 0.5 mm (for neonatal rats) or 0.7 mm (for young/adult rats). 13b. Using a programmable syringe pump, inject the cell suspension at a rate of 0.200 μl/min. 14b. Leave the micropipet in the spinal cord for 5 min after injection (to prevent the backflow of cells), then slowly withdraw the micropipet.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

B104 collection medium (500 ml) 487.65 ml DMEM (Invitrogen, cat. no. 12100-046) + 3.7 g/liter NaHCO3 (Sigma, cat. no. S-5761) 2.5 ml of 200× N1 supplement (see recipe) 250 μl of 10 mg/ml insulin (Sigma, cat. no. I-6634) in 0.01 N HCl 4.6 ml of 100 mM sodium pyruvate (Sigma, cat. no. 8636) 5.0 ml of 100× penicillin/streptomycin (Invitrogen, cat. no. 15140-122; 1× final concentration) Store up to 1 month at 4◦ C B104 feeding medium (500 ml) 440.4 ml DMEM (Invitrogen, cat. no. 12100-046) + 3.7 g/liter NaHCO3 (Sigma, cat. no. S-5761) 50 ml fetal bovine serum (FBS; Hyclone, cat. no. SH30070.03) 4.6 ml of 100 mM sodium pyruvate (Sigma, cat. no. S8636) 5.0 ml of 100× penicillin/streptomycin (Invitrogen, cat. no. 15140-122; 1× final concentration) Store up to 1 month at 4◦ C DMEM/F12, 10× (200 ml) Add 1 packet DMEM (Invitrogen, cat. no. 12100-046) and 1 packet F12 (Sigma, cat. no. N-6760) to 100 ml ddH2 O and mix. Bring volume to 200 ml and filtersterilize through a 0.22-μm filter (Millipore, cat. no. SCGPU05RE). Store up to 3 months at 4◦ C.

N1 supplement, 200× 80.55 mg putrescine (Sigma, cat. no. P-7505) 20 ml Hanks’ Balanced Salt Solution (HBSS; Invitrogen, cat. no. 14175) 50 μl of progesterone (Sigma, cat. no. P-9783), 2 mM in ethanol 50 ml of sodium selenite (Sigma, cat. no. S-5261), 3 mM in ddH2 O 5 ml of 5 mg/ ml apo-transferrin (Sigma, cat. no. T-2036) in PBS Filter-sterilize through a 0.22-μm filter (Millipore, cat. no. SCGPU05RE) Store 1-ml aliquots up to 12 months at −20◦ C Neurosphere medium (500 ml) Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

358.75 ml sterile ddH2 O 50 ml of 10× DMEM/F12 (see recipe) 10 ml of 30% (w/v) glucose (Sigma, cat. no. G-7021) 7.5 ml of 7.5% (w/v) NaHCO3 (Sigma, cat. no. S-5761) 2.5 ml of 1 M HEPES (Sigma, cat. no. H-0887) continued

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1.25 ml of 4% (w/v) BSA in HBSS (Sigma, cat. no. A-7906) 10 ml of 100 mg/ml heparin (Sigma, cat. no. H-3149) in 1× DMEM/F12 5 ml of 100× L-glutamine (Invitrogen, cat. no. 25030-081; 1× final concentration) 5 ml of 100× penicillin/streptomycin (Invitrogen, cat. no. 15140-122; 1× final concentration) 50 ml of 10× neurosphere medium hormone mix (see recipe) Store up to 1 month at 4◦ C Important Note: BSA must be added prior to the hormone mix to prevent precipitation.

Neurosphere medium hormone mix, 10× 100 ml of 10× DMEM/F12 (see recipe) 20 ml of 30% (w/v) glucose (Sigma, cat. no. G-7021) 15 ml of 7.5% (w/v) NaHCO3 (Sigma, cat. no. S-5761) 5 ml of 1 M HEPES 750 ml sterile ddH2 O 100 mg Apo-transferrin (Sigma, cat. no. T-2036) 100 ml of 2.5 mg/ml insulin (Sigma, cat. no. I-6634) in 0.01 N HCl 6 ml of 0.966 mg/ml putrescine (Sigma, cat. no. P-7505) in ddH2 O 100 μl sodium selenite (Sigma, cat. no. S-5261), 3 mM in ddH2 O 100 μl progesterone (Sigma, cat. no. P-9783), 2 mM in ethanol Store 25-ml aliquots up to 12 months at –20◦ C Oligosphere differentiation medium (100 ml) 98.85 ml DMEM (Invitrogen, cat. no. 12100-46) 500 μl of 200× N1 supplement (see recipe) 500 μl fetal bovine serum (FBS; Hyclone, cat. no. SH30070.03) 50 μl of 10 mg/ml insulin (Sigma, cat. no. I-6634) in 0.01 N HCl 100 μl of 10 μg/ml biotin (Sigma, cat. no. B-4501) 1 ml of 100× penicillin/streptomycin (Invitrogen, cat. no. 15140-122; 1× final concentration) Store up to 1 month at 4◦ C Oligosphere medium (100 ml) 70 ml neurosphere medium (see recipe) 30 ml B104-conditioned medium (see recipe) Store up to 1 month at 4◦ C Pulled glass micropipets Pull a borosilicate glass capillary (World Precision Instruments, cat. no. 1B100F-4) with a needle/pipet puller (David Kopf Instruments, cat. no. 720) to make a micropipet. Connect the micropipet with a Hamilton gastight syringe (10 μl; Hamilton, cat. no. 1701) with PTFE tubing.

COMMENTARY Background Information Avellana-Adalid et al. (1996) first described a technique for the culturing and expansion of free-floating oligodendrocyte progenitor cells from the rat neonatal brain. The aggregates of oligodendrocyte progenitors produced by this technique were dubbed “oligospheres” by the authors. The authors further demonstrated that these OPCs could be differentiated into oligodendrocytes and were capable of myeli-

nation following transplantation into the brain of newborn shiverer mice. While the techniques described in this paper have since been modified, the use of conditioned medium from the B104 neuroblastoma cell line in the generation of oligospheres has been a constant. OPCs may potentially be used as an exogenous source of cells to treat lesions in multiple sclerosis (Duncan, 2008) and genetic myelin disorders (Duncan, 2005). However, the

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validity of this approach must first be demonstrated in animal models prior to clinical trials in humans. Rat OPCs have a number of advantages over OPCs from other sources, such as mice. Many of the models for genetic myelin disorders have either been discovered in rats, such as the myelin-deficient (md) rat (Csiza and de Lahunta, 1979) and the Long Evans Shaker (les) rat (Delaney et al., 1995), or generated by transgenic approaches, such as the PLP-overexpressing rat (Bradl et al., 1999). Likewise, rats immunized with fragments of myelin proteins develop experimental autoimmune encephalomyelitis (EAE), a commonly used animal model for multiple sclerosis (Gold et al., 2006). Transplanting wild-type rat OPCs into rat myelin disease models or rat EAE models eliminates the need for immunosuppressive drugs to prevent rejection of transplanted cells. Furthermore, oligosphere cultures from murine or canine sources exhibit slowed growth after 2 months, and cells that migrate out from these spheres are poor sources of myelinating cells for transplantation (Zhang et al., 1998b). In contrast, rat oligosphere cultures exhibit no growth deficiencies and produce excellent cells for transplantation up to 6 months after culturing.

Critical Parameters and Troubleshooting

Generation of Cultured Oligodendrocyte Progenitor Cells from Rat Neonatal Brains

In this unit we have described the protocol for isolating OPCs from rats in a post-natal age range of 0 to 6 days. In our past experience we have found that the younger the animal, the more robust and longer-lived the cultured OPCs. Typically, we prefer pups at a post-natal age of approximately day 2, as the dissection is relatively easy and cultured OPCs derived from these pups will survive longer than OPCs from older neonates. After a post-natal age of 6 days, the number of OPCs that can be isolated from rat striatum drops significantly, as more and more of these cells will have differentiated into oligospheres and other glial cell types. OPC transplantation into rat models of myelin disease, such as the md and les rats, is a commonly used model system. However, given the short life span of the md rat (∼21 days), OPC transplantation into neonatal pups should be performed by post-natal day 7 to provide transplanted cells sufficient time to differentiate and myelinate. In the les rat, OPCs should likewise be transplanted into the spinal cord by post-natal day 7; after this time-point, an increase in microglial activation results in greatly reduced survival of transplanted OPCs (Zhang et al., 2003).

Anticipated Results The culture protocol outlined in this unit can be expected to generate a highly purified population of oligodendrocyte progenitor cells from the rat neonatal brain. Directly culturing rat striatal tissue in B104-conditioned medium, or switching neurospheres into this medium, should yield a population of ∼100% OPCs (Zhang et al., 1998a). Following exposure to differentiation medium, rat oligospheres will differentiate to a population of cells consisting of >95% oligodendrocytes, as identified by positive staining for O4 and MBP, among other oligodendrocyte markers (Zhang et al., 1998a). The remaining cells will be positive for GFAP, identifying them as astrocytes. However, the ratio of oligodendrocytes to astrocytes produced following OPC differentiation will be dependent on a number of factors, such as the species that acts as a source of OPCs, the age of the animal used to generate the oligospheres, and the length of time the oligospheres have been in culture. Rat OPCs typically have a higher percentage of progenitors differentiating into oligodendrocytes than OPCs from other species. Similarly, the percentage of OPCs that will differentiate into oligodendrocytes following transplantation into a rat EAE or myelin disease model will be dependent not only on the nature of the cells being transplanted, but also on the environment that they are being transplanted into. That said, transplantation of rat OPCs into the brain and spinal cord of the md rat is predicted to result in widespread migration of progenitors through the white matter and extensive myelination that can be observed 2 weeks after transplantation (Tontsch et al., 1994; Learish et al., 1999). Similar results can be expected when OPCs are transplanted into the les rat, provided the transplantation is performed prior to microglial activation (Zhang et al., 2003).

Time Considerations

It will take ∼4 weeks in the presence of oligosphere medium for rat neonate striatal cultures to contain primarily oligospheres. Oligospheres will continue to proliferate for many months after culturing. However, with increasing age (>6 months), the rate of oligosphere proliferation will decline. Furthermore, following exposure to differentiation medium, a greater number of these oligospheres will differentiate into astrocytes, which can be identified by GFAP immunolabeling. Therefore, it is advisable to either use oligospheres within the first few months after the generation of OPC

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cultures, or store them in liquid nitrogen for later usage. The transition of oligospheres to oligodendrocytes following exposure to differentiation medium is a process that can be followed both by observation of morphological changes and by immunostaining for immature and mature oligodendrocyte markers. After 1 to 3 days of exposure to oligosphere differentiation medium, O4 staining will be visible. After ∼1 week in differentiation medium, staining for the myelin markers PLP and MBP will begin to be visible. After 2 weeks in differentiation medium, differentiation into immature and mature oligodendrocytes will largely be complete.

Acknowledgement This work was supported by a grant from the National Multiple Sclerosis Society (no. TR3761).

Literature Cited Asakura, K., Hunter, S.F., and Rodriguez, M. 1997. Effects of transforming growth factor β and platelet-derived growth factor on oligodendrocyte precursors: Insights gained from a neuronal cell line. J. Neurochem. 68:2281-2290. Avellana-Adalid., V., Nait-Oumesmar, B., Lachapelle, F., and Baron-Van Evercooren, A. 1996. Expansion of rat oligodendrocyte progenitors into proliferative “oligospheres” that retain differentiation potential. J. Neurosci. Res. 45:558-570. Bradl, M., Bauer, J., Inomata, T., Zielasek, J., Nave, K.-A., Toyka, K., Lassmann, H., and Wekerle, H. 1999. Transgenic Lewis rats overexpressing the proteolipid protein gene: Myelin degeneration and its effect on T cell-mediated experimental autoimmune encephalomyelitis. Acta Neuropathol. 97:595-606. Csiza, C.K. and de Lahunta, A. 1979. Myelin deficiency (md): A neurologic mutant in the Wistar rat. Am. J. Pathol. 95:215-223.

Delaney, K.H., Kwiecien, J.M., Wegiel, J., Wisniewski, H.M., Percy, D.H., and Fletch, A.L. 1995. Familial dysmyelination in a Long Evans rat mutant. Lab. Anim. Sci. 45:547553. Duncan, I.D. 2005. Oligodendrocytes and stem cell transplantation: Their potential in the treatment of leukoencephalopathies. J. Inherit. Metab. Dis. 28:357-368. Duncan, I.D. 2008. Replacing cells in multiple sclerosis. J. Neurol. Sci. 265:89-92. Gold, R., Linington, C., and Lassmann, H. 2006. Understanding pathogenesis and therapy of multiple sclerosis via animal models: 70 years of merits and culprits in experimental autoimmune encephalomyelitis research. Brain 129:19531971. Learish, R.D., Brustle, O., Zhang, S.-C., and Duncan, I.D. 1999. Intraventricular transplantation of oligodendrocyte progenitors into a fetal myelin mutant results in widespread formation of myelin. Ann. Neurol. 46:716-722. Tontsch, U., Archer, D.R., Dubois-Dalcq, M., and Duncan, I.D. 1994. Transplantation of an oligodendrocyte cell line leading to extensive myelination. Proc. Natl. Acad. Sci. U.S.A. 91:1161611620. Utzschneider, D.A., Archer, D.R., Kocsis, J.D., Waxman, S.G., and Duncan, I.D. 1994. Transplantation of glial cells enhances action potential conduction of amyelinated spinal cord axons in the myelin-deficient rat. Proc. Natl. Acad. Sci. U.S.A. 91:53-57. Zhang, S.-C., Lundberg, C., Lipsitz, D., O’Connor, L.T., and Duncan, I.D. 1998a. Generation of oligodendroglial progenitors from neural stem cells. J. Neurocytol. 27:475-489. Zhang, S.-C., Lipsitx, D., and Duncan, I.D. 1998b. Self-renewing canine oligodendroglial progenitors expanded as oligospheres. J. Neurosci. Res. 54:181-190. Zhang, S.-C., Goetz, B.D., and Duncan, I.D. 2003. Suppression of activated microglia promotes survival and function of transplanted oligodendroglial progenitors. Glia 41:191-198.

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Isolating, Expanding, and Infecting Human and Rodent Fetal Neural Progenitor Cells

UNIT 2D.2

Allison D. Ebert,1 Erin L. McMillan,1 and Clive N. Svendsen1, 2 1 2

Waisman Center, University of Wisconsin-Madison, Madison, Wisconsin Department of Anatomy, University of Wisconsin-Madison, Madison, Wisconsin

ABSTRACT Neural progenitor cells have tremendous utility for understanding basic developmental processes, disease modeling, and therapeutic intervention. The protocols described in this unit provide detailed information to isolate and expand human and rodent neural progenitor cells in culture for several months as floating aggregates (termed neurospheres) or plated cultures. Detailed protocols for cryopreservation, neural differentiation, exogenous gene expression using lentivirus, and transplantation into the rodent nervous C 2008 by system are also described. Curr. Protoc. Stem Cell Biol. 6:2D.2.1-2D.2.16.  John Wiley & Sons, Inc. Keywords: stem cells r brain r in vitro r mouse r rat r embryonic r neural progenitor cells

INTRODUCTION Neural progenitor cells (NPCs) are stem cells whose lineage potential has been restricted to solely the central nervous system (CNS). They have tremendous utility for understanding basic developmental processes, disease modeling, and therapeutic intervention. The protocols provided in this unit describe the procedures relating to the growth and maintenance of human and rodent neural progenitor cells from fetal tissues. Adult neural progenitor cells are discussed in other protocols, so they will not be addressed here. Each of the five protocols addresses a different aspect of the growth and propagation of these cells. The first protocol provides the steps to isolate neural progenitor cells from primary human, rat, or mouse fetal tissue (Basic Protocol 1) and propagate them as neurospheres (i.e., floating aggregates) for many weeks. Alternate Protocol 1 describes a procedure for culturing neural progenitor cells as single cells. Growth and expansion of NPCs by enzymatic digestion is described in Basic Protocol 2, and growth and expansion of these cells by mechanical chopping is described in Alternate Protocol 2. Support Protocol 1 provides details for clonal analysis of the cells. Basic Protocol 3 provides the steps to completely dissociate human and rodent cells in preparation for plating onto a permissive substrate to promote differentiation into post-mitotic neural cells (specifically neurons and astrocytes), or for transplantation into the rodent central nervous system. Basic Protocol 4 outlines the steps necessary to infect progenitor cells with a lentivirus to force overexpression of various transgenes; Alternate Protocol 3 describes these methods for fetal rodent cells. This is a useful technique to test the influence of various transcription factors on growth and differentiation, and can also be used to generate stable lines of protein- or drug-secreting cells. Basic Protocol 5 explains the procedures to cryopreserve the progenitor cells for long-term storage and subsequently thaw them for later use. The ability of the progenitor cells to be frozen and thawed increases their utility for various in vitro and in vivo experiments. There is also a protocol for coating coverslips for culture (Support Protocol 2). Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.2.1-2D.2.16 Published online September 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d02s6 C 2008 John Wiley & Sons, Inc. Copyright 

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There are hundreds of published papers using a wide variety of methods for growing neural progenitor cells. We have attempted to describe a “generic” set of protocols which are to some degree based around our own experience with these cells over the past 20 years. However, we expect users to optimize these fundamental protocols based on the extensive literature on this topic. CAUTION: Primary human tissue, human progenitor cell cultures, and any medium removed from these cultures are hazardous waste and should be contained and discarded in appropriate biohazard containers. NOTE: All procedures should be completed in a laminar-flow culture hood unless indicated otherwise. When transferring samples to the water bath or incubator, make sure all lids are on and closed tightly. All cultures are maintained in a humidified incubator at 37◦ C and 5% CO2 and all media are warmed in a 37◦ C water bath prior to use. NOTE: As stated in the “Guidelines for the Conduct of Human Embryonic Stem Cell Research” (see APPENDIX 1A), human tissue research must be reviewed and approved by the institutional ethics review panel, and donated material must be provided voluntarily with informed consent. NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to government regulations for the care and use of laboratory animals. BASIC PROTOCOL 1

ISOLATING NEURAL PROGENITOR CELLS FROM HUMAN AND RODENT TISSUE AND NEUROSPHERE CULTURE The protocol outlined below explains how to dissect tissue from any region of the developing embryo to generate neural progenitor cell cultures and propagate them as floating neurospheres, as previously identified by Reynolds and Weiss (1992). Depending on the region, age, and species of fetal tissue used, the growth rates and differentiation properties will vary (Svendsen et al., 1997; Laywell et al., 2000; Hitoshi et al., 2002; Ostenfeld et al., 2002; Watanabe et al., 2004; Kim et al., 2006). When cells are grown as neurospheres in the presence of mitogens (EGF, FGF), they are maintained in the undifferentiated/uncommitted state. Cells at this stage will be >90% positive for nestin, but within neurospheres there may be a complex mix of stem cells and progenitors at various stages of differentiation. This is somewhat dependant on the size of the neurospheres, because as they get larger there is more chance for differentiation. Once neurospheres are removed from these mitogens and plated on a permissive substrate (see Basic Protocol 3), they will adopt characteristics of terminally differentiated neural cells (e.g., βIII-tubulin positive neurons and GFAP positive astrocytes). Alternative growth and propagation methods as plated cells are also included.

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Materials Human fetal tissue or rat embryos of the appropriate age (e.g., ED 15) 0.6% (w/v) glucose in PBS without Ca or Mg (Invitrogen, cat. no. 14190-250) 0.05% (w/v) trypsin/EDTA (Invitrogen) 1× soybean trypsin inhibitor (can be purchased from various companies) 1 U/μl DNase I (3360 U/mg, Sigma) in 0.6% glucose/PBS Starting medium (see recipe) Maintenance medium (see recipe) 70% ethanol Laminar-flow hood with microscope Dissecting tools: Microscissors Sharp forceps (5 point) Current Protocols in Stem Cell Biology

10-cm culture dishes Hemacytometer (also see UNIT 1C.3) 25-cm2 , 75-cm2 , and/or 175-cm2 filter top culture flasks (will vary) Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) Collect tissue 1. Obtain human fetal tissue in 0.6% glucose in PBS or isolate rodent embryos at the appropriate gestational age (e.g., embryonic day 15). Place the tissue in a 10-cm culture dish containing cold PBS and gently agitate the dish to wash the tissue. In general, combine tissue from five to eight rodent embryos to make one culture. Human fetal tissue may be difficult to obtain in quantities larger than 1 mg, so combine all available tissue into one culture. The gestational age required for both rodent and human tissue will depend on the region being collected (e.g., embryonic day 15 for mouse striatum).

2. Working in a laminar-flow hood, microdissect the tissue of interest (e.g., cortex, ventral mesencephalon, hippocampus, spinal cord) under a dissecting microscope in a series of 10-cm culture dishes containing ice-cold 0.6% glucose in PBS. Keep moving the tissue of interest to new dishes to isolate it from the discarded tissue. If dissections cannot be performed in a laminar-flow hood, be careful not to contaminate the tissues. Use autoclaved dissecting tools and clean the microscope and work area with 70% ethanol before starting. There is no specific size of tissue that is desirable, because that will vary depending on the region of interest. When dissecting the same region from multiple embryos, it is ideal to have all the tissue pieces dissected in the same way, which would give tissue pieces of the same size.

Dissociate tissue 3. Put dissected pieces into a sterile microcentrifuge tube and add 1 ml of 0.05% trypsin/EDTA. Incubate 10 to 20 min in a 37◦ C water bath. Do not centrifuge between steps 3 and 5; just allow tissue to settle by gravity. From this point on, the tissue should only be handled inside a laminar-flow culture hood.

4. Remove as much trypsin as possible, replace with 1 ml of 1× trypsin inhibitor, and incubate for 10 min in a 37◦ C water bath. 5. Remove trypsin inhibitor, replace with 1 ml of 1 U/μl DNase I, and incubate for 10 min in a 37◦ C water bath. 6. Remove DNase I and replace with 1.0 ml starting medium. 7. Triturate by passing through a 1000-μl pipet tip and then through a 200-μl pipet tip, until a single-cell suspension is obtained. Pass through each tip ∼20 times.

Plate cells 8. Using a hemacytometer, count cells in a 10-μl aliquot and assess viability using 0.4% trypan blue (UNIT 1C.3). Any dilution is acceptable for counting. One example would be to add 10 μl of cell suspension to 90 μl of medium and mix well. Add 50 μl of this 10× dilution to 50 μl of the prepared trypan blue solution and load the hemacytometer with 10 μl of this solution. The total sample dilution is 20×. Count five squares on the hemacytometer and determine the average number of cells. Multiply this value by the dilution factor and then by 10,000 to give number of cells/ml. Make adjustments for the actual volume of the cell suspension to calculate the total number of cells available.

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9. Seed the cells into the appropriate-sized flask at a density of 200,000 cells/ml in starting medium. Many neural progenitor cell protocols use B27 supplement in the starting medium, as it provides antioxidant support during the initial stress of plating and increases growth and survival (Svendsen et al., 1995). In general, use 10 ml total medium in a 25-cm2 flask, 20 ml total in a 75-cm2 flask, or 40 ml total in a 175-cm2 flask. Cells should form spheres within 2 to 3 days.

10. Feed flasks every 3 or 4 days by allowing neurospheres to settle in each flask and then removing half the conditioned medium and replacing it with fresh medium. Be careful not to discard any spheres. Spheres are maintained in starting medium during the initial growth period (∼1 week for rodent and 4 weeks for human tissues) and passaged (see Basic Protocol 2) generally every 7 to 10 days. After the initial growth period, neurospheres can be switched to maintenance medium. After approximately 10 passages, leukemia inhibitory factor (LIF) should be added to the maintenance medium to extend neurosphere expansion.

11. Clean all work areas with 70% ethanol and dispose of tissue, plates, and discarded solutions as biohazard waste. Each human tissue sample or group of rodent embryos should be considered an independent line and should not be combined with other samples/lines. ALTERNATE PROTOCOL 1

BASIC PROTOCOL 2

CULTURING NEURAL PROGENITOR CELLS AS SINGLE CELLS Rather than growing the cells as neurospheres, fresh progenitor isolates can be plated as single cells on poly-ornithine (15 μg/ml) and laminin (5 to 10 μg/ml)– or fibronectin (1 μg/ml)–coated tissue culture flasks or plates using either starting or maintenance medium (Ray et al., 1993; Ray and Gage, 1994; Johe et al., 1996). See Support Protocol 2 for the coating of culture plates, flasks, and coverslips. However, a recent report suggests mouse whole-brain progenitor cells grow well on uncoated culture plates (Ray and Gage, 2006). Coating culture plates or flasks takes at least overnight for the poly-ornithine and can require another night for the laminin or fibronectin depending on the cell source used and user preference. The cells can be passaged when confluent using standard enzymatic/mechanical dissociation (see Basic Protocol 2). The density of reseeding can vary, but can range from 20,000 to 45,000 cells/cm2 .

GROWING AND EXPANDING NEURAL PROGENITOR CELLS BY ENZYMATIC DISSOCIATION There are two basic methods for passaging neurospheres, enzymatic dissociation (this protocol) or chopping (Alternate Protocol 2). Confluent plated cultures require enzymatic digestion, lifting, and reseeding. Floating neurospheres are normally passaged approximately every 7 days when they reach 400 to 500 μm in diameter. Cells should re-form spheres within 2 to 3 days after enzymatic dissociation or chopping.

Materials Neurospheres (Basic Protocol 1)w 0.05% (w/v) trypsin/EDTA (Invitrogen) 1× soybean trypsin inhibitor (can be purchased from various companies) Base medium (see recipe) Plating medium (see recipe) Starting/maintenance medium (see recipe) Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

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15-ml conical centrifuge tubes Hemacytometer (also see UNIT 1C.3) 25-cm2 , 75-cm2 , and/or 175-cm2 filter top culture flasks (will vary) Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) Current Protocols in Stem Cell Biology

1. Allow neurospheres to settle in the flask and carefully transfer the spheres into a 15-ml conical tube. Be sure to settle and collect any spheres adhering to the bottom and sides of the flask.

2. Remove and discard the medium, add 1 ml of 0.05% trypsin/EDTA, and incubate for 10 min in a 37◦ C water bath. 3. Remove the trypsin, replace with 1 ml of 1× trypsin inhibitor, and incubate for 10 min in a 37◦ C water bath. 4. Remove the trypsin inhibitor and replace with 10 ml base medium. Gently mix the cells and allow cells to settle by gravity. 5. Remove the base medium and replace with 1 ml plating medium. Starting or maintenance media are acceptable in place of plating medium; however, using plating medium does not consume costly mitogens (EGF and FGF).

6. Triturate cells by passing through a 200-μl pipet tip (40 to 50 times) to make a single-cell suspension. A glass Pasteur pipet can also be used.

7. Using a hemacytometer, count cells and assess viability using trypan blue (UNIT 1C.3) in an aliquot of cell suspension. Follow a similar dilution as described in the annotation to step 8 of Basic Protocol 1.

8. Seed the cells into the appropriate sized flask at a density of 100,000 cells/ml in starting/maintenance medium. 9. Feed flasks every 3 or 4 days by settling cells in flask, removing all of the medium, and replacing it with fresh starting/maintenance medium. Our experience is that while mouse neurospheres will continue to grow for many weeks or months, rat neurospheres will undergo senescence within 6 weeks (Svendsen et al., 1997). While the reason for this remains unclear, mouse cells are prone to genomic instability (Todaro and Green, 1963) and may transform in vitro at later passages (Morshead et al., 2002). Rat neurospheres are less likely to transform and follow a senescence pattern in vitro. Human neurospheres generated from the cortex can grow for approximately 50 weeks before senescing (Wright et al., 2006).

GROWING AND EXPANDING NEURAL PROGENITOR CELLS BY MECHANICAL CHOPPING

ALTERNATE PROTOCOL 2

While rodent cultures are easy to expand using any passaging method, human neural progenitor cells do not grow as fast and can be difficult to maintain. There are various ways to grow human neural progenitor cells, but one simple method to increase growth and survival is to use a nonenzymatic, mechanical passaging method (Svendsen et al., 1998). This method also works in the absence of FGF-2, which can be expensive to add to the medium of bulk cultures over long time periods. Cultures should be chopped when the majority of the spheres are approximately 400 to 500 μm in diameter (generally every 7 to 10 days). This chopping method does not allow for clonal analysis. If dissociation and passaging of the cells is required (e.g., for clonal analysis; see Support Protocol 1), FGF-2 and heparin should be added to the maintenance medium to help with the survival and growth of the cultures.

Materials Human neurospheres (Basic Protocol 1) 70% and 100% ethanol Starting and/or maintenance medium (depending on the age or passage number; see recipes and annotation to step 10 of Basic Protocol 1)

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McIlwain tissue chopper (Lafayette Instruments, model no. TC752) Blunt forceps Small beaker Bead sterilizer (optional; e.g., WU-10779-00, Cole-Parmer) Double-edged razor blade (e.g., Fisher, cat. no. NC9732480) 15- and 50-ml conical centrifuge tubes Narrow profile 50-mm culture dish (will only use the lid) 25-cm2 , 75-cm2 , and/or 175-cm2 filter-top culture flasks (as needed for expanding cell number) Prepare for passaging of human neurospheres 1. Put the McIlwain tissue chopper into a laminar flow culture hood and clean with 70% ethanol. 2. Using forceps, soak a double-edged razor blade in 100% ethanol (in the small beaker) and flame sterilize. Some institutions discourage or prohibit open flames in the culture hoods, so heating the razor blade in a dry glass-bead sterilizer is acceptable.

3. Carefully secure the blade onto the chopping arm. With the chopping arm in the highest position, remove the screw and plate, insert the blade, and slightly tighten the screw to fix the plate over the blade. Make sure the blade is parallel to the chopping surface by carefully adjusting the blade and tightening the screw.

4. Check the chopper settings, turn on the power, and press “reset.” The blade force control should be set at 12:00 (straight up), and, for an optimal chop, the chop distance should be set at 200 μm. Ensure that there is enough vacuum grease on the base of the chopping disc to allow for smooth plate movement.

5. Choose the appropriate number and size of flasks needed for the newly chopped cultures. Depending on the size and density of the spheres to be chopped, cultures can be split into multiple flasks (e.g., one into two, one into three), or can be put into a larger flask (e.g., a 25-cm2 into a 75-cm2 ). Keep in mind, however, that cells generally recover from the chop better in a slightly more populated culture environment.

Chop the neurospheres Sister cultures can be pooled for chopping, but do not pool more than two 175-cm2 flasks for a single chop. 6. Settle neurospheres in the flask. Lean the flask so that it is resting on its bottom corner. Be sure to settle and collect any spheres adhering to the bottom and sides of the flask.

7. Once spheres are settled, remove a majority of the conditioned medium (CM) and transfer to a 50-ml conical tube for later use. 8. Transfer the neurospheres and remaining medium to a 15-ml conical tube and allow the spheres to settle. Do not centrifuge; allow spheres to settle by gravity. Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

9. Once settled, use a glass Pasteur pipet to transfer the spheres to the middle of the inverted lid of 50-mm culture dish. 10. Carefully remove as much of the medium as possible with a Pasteur pipet, leaving the spheres in the center of the lid, and place the medium back into the 15-ml conical tube (from step 8).

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It is important to remove as much medium as possible in order to prevent the spheres from moving during the chop. Removing the medium without removing spheres can be difficult, but tilting the lid generally helps pool the medium to one side leaving a cluster of spheres in the center of the lid. Inevitably, there will be some spheres that are removed with the medium. Transfer these back into the 15-ml conical tube. Preferably, and if many large spheres have been removed, let them settle, and put them back on the plate to try again. Otherwise, leave them in the 15-ml conical tube and move on.

11. Place the dish lid on the chopper and move the sliding table to the starting position. The chopping table moves from left to right, so the cells should be in the far-left position with the chopping arm and blade to the right.

12. Start chopping by slowly turning the speed dial clockwise to the 12:00 position. The blade will leave lines on the plastic lid. If the blade is not aligned parallel to the chopping surface, the lines will be noticeably uneven. Turn the speed dial down to stop the chopping arm and carefully readjust the blade.

13. When all of the spheres on the lid have been chopped, stop and raise the chopping arm, and reposition the table to the starting position. 14. Rotate the dish lid 90◦ and repeat steps 12 and 13 one more time.

Collect the chopped spheres 15. When the spheres have been chopped in the second direction, remove the dish from the chopper. 16. Add a small amount of CM (from step 7) to the cells and transfer them using a Pasteur pipet back into the 15-ml conical tube used in step 8. In order to remove the cells from the dish after chopping, it may be necessary to gently scrape the plate with the tip of the Pasteur pipet and repeatedly wash the plate with conditioned medium. Some slivers of plastic may get into the culture, but these do not seem to harm the spheres or hinder their growth.

17. Add more CM (from step 7) to the 15-ml conical tube and gently resuspend with a 10-ml pipet. 18. Evenly distribute the cell suspension to the new flasks. Keep in mind that half of the total medium volume in the flask should be fresh medium and half should be CM. For example, in a 75-cm2 flask, add 10 ml CM and 10 ml fresh maintenance medium. Use the CM collected in step 7 to divide among the new flasks. Do not cross-contaminate cultures with CM. Pooling CM from sister cultures is acceptable, but refrain from using CM from different lines.

19. Repeat steps 2 to 18 for additional cultures to chop. To prevent cross-contamination, clean the chopper with 70% ethanol and UV irradiate everything in the culture hood between chops of different lines. If multiple flasks of sister cultures are being chopped, it is not necessary to UV sterilize between chops, but use a new, sterilized blade.

20. After all chopping is complete, extinguish the flame, discard the razor blade, and dispose of all plates and used medium as biohazard waste. Clean the chopper and all work areas with 70% ethanol.

CLONAL ANALYSIS OF NEUROSPHERES To test the clonality of both rodent and human cells that are capable of forming spheres (Reynolds and Weiss, 1996; Vescovi et al., 1999), neurospheres are completely dissociated (Basic Protocol 2) and serially diluted to ∼1 to 2 cells/10 μl in starting medium (see recipe) in a final volume sufficient to seed multiple wells. Ten μl of the cell suspension is

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plated into each well of a 96-well plate. Inspect the wells 24 hr after plating to ensure that only single cells are in each well. After 7 to 10 days, inspect the wells for the presence or absence of spheres. FGF-2 (20 ng/ml) and heparin (5 μg/ml) should be added to the maintenance medium (see recipe) in order to promote the growth of cells under these cloning conditions. The efficiency of this method is often very low (<5%), and in our experience, it is very difficult to achieve clonal analysis with human neural progenitor cells. BASIC PROTOCOL 3

COMPLETE DISSOCIATION OF NEURAL PROGENITOR CELLS FOR TERMINAL DIFFERENTIATION AND/OR TRANSPLANTATION Human and rodent neural progenitor cells terminally differentiate into neurons, astrocytes, and oligodendrocytes when provided with the appropriate substrate (laminin) and grown in the absence of mitogens (EGF, FGF-2). In the authors’ experience, oligodendrocytes are only seen in very low numbers from human fetal brain cultures grown as neurospheres at early passages, and these are reduced further upon expansion (Chandran et al., 2004). However, other methods of growing human progenitors are being developed (e.g., on laminin) that may improve oligodendrocyte production. Rodent neurospheres from the brain and spinal cord give rise to many oligodendrocytes at all passages (Ostenfeld et al., 2002; Chandran et al., 2004).

Materials Neurospheres (Basic Protocol 1) 1× Accutase (Millipore, cat. no. SCR005, also available from Sigma; for human cells) or 0.05% trypsin/EDTA (e.g., Invitrogen; rodent cells) 1× trypsin inhibitor (e.g., Invitrogen) Base medium (see recipe) 50 μg/ml laminin (mouse; available from many vendors) Dulbecco’s Modified Eagle Medium (DMEM, e.g., Invitrogen) Plating medium (see recipe) Transplant medium (see recipe) 15-ml conical tubes Hemacytometer (also see UNIT 1C.3) Bellco round glass coverslips (Fisher), poly-ornithine-coated 24-well tissue culture plate Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) Dissociate neural progenitor cells 1. Allow neurospheres to settle in the flask and carefully transfer the spheres into a 15-ml conical tube. Be sure to settle and collect any spheres adhering to the bottom and sides of the flask. Generally, dissociation of human cells is done 3 to 4 days after chopping.

2. Remove and discard the medium, replace with 1 ml of 1× Accutase for human cells or 1 ml 0.05% trypsin/EDTA for rodent cells, and incubate for 10 min in a 37◦ C water bath.

Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

The Accutase has less adverse effect on the overall health of the human cells than trypsin. Keep the Accutase frozen until ready to use and then warm an aliquot for 5 min in a 37◦ C water bath. Gently swirl the cells once during the incubation.

3. Remove the Accutase or trypsin, replace with 1 ml of 1× trypsin inhibitor (for both human and rodent cultures), and incubate for 10 min in a 37◦ C water bath.

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4. Remove the trypsin inhibitor and add 10 ml base medium. Gently mix the cells and allow cells to settle by gravity.

5. Remove the base medium and replace with 1 ml plating medium. 6. Triturate cells by passing through a 200 μl pipet tip (40 to 50 times) to make a single-cell suspension. 7. Using a hemacytometer, count an aliquot of cells and assess viability using trypan blue (UNIT 1C.3) in an aliquot of cells. If differentiating the cells, go to step 8; if transplanting the cells, go to step 13. Follow a similar dilution as described in step 8 of Basic Protocol 1.

Plate progenitor cells for differentiation 8. Place a poly-ornithine-coated coverslip in each well of a 24-well tissue culture plate under a laminar-flow culture hood. Apply 30 μl laminin to the center of each poly-ornithine-coated coverslip and incubate for 30 min at 37◦ C. 9. Remove and discard laminin and wash three times with 30 to 50 μl of DMEM. Do not flood the wells with DMEM because only 30 μl of cells will be plated (see below), and it is important that the cells stick to the laminin spot. It is also important to wait to remove the last DMEM wash until ready to plate the cells on the coverslip. This prevents excessive drying of the laminin.

10. Plate 30,000 cells total on each coverslip by adjusting the volume of plating medium to obtain a final concentration of 1000 cells/μl and placing 30 μl of cell suspension on each coverslip. 11. Place the plate into the incubator for 1 hr to allow the cells to attach to the coverslip, then flood each well with 1 ml plating medium. 12. Replace half of the medium with fresh plating medium every 3 to 4 days for 1 to 2 weeks. Cells differentiate in the absence of EGF and FGF, so use only plating medium. Cells will differentiate during the first week, but 1% (v/v) FBS may be added to the cultures to aid survival for a full 2 weeks. Cells can then be processed and analyzed as desired (e.g., by immunocytochemistry or immunoblotting).

Collect cells for transplantation 13. Calculate the number of cells necessary for the transplant experiments. The optimal number of cells to transplant will vary depending on the experimental design—e.g., 1 × 106 total cells injected into a rat striatum or 3 × 105 total cells injected into the mouse striatum. The volume injected will vary, but a typical range would be from 1 to 10 μl.

14. Transfer the 1 ml of the cell suspension (from step 6) into a 1.5-ml microcentrifuge tube. 15. Microcentrifuge 1 to 2 min at 2000 rpm, room temperature, to pellet cells. 16. Remove all the medium and resuspend in the appropriate volume of transplant medium to obtain the proper final cell concentration as determined in step 12. 17. Place the cells on ice and keep on ice throughout the transplantation procedure. 18. Using a hemacytometer, recount the an aliquot of cells and assess viability using trypan blue (UNIT 1C.3). It is important to repeat the cell count at this time to verify the cell concentration prior to transplantation.

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19. To assess the viability of any extra cells dissociated for transplantation, but not used, follow the “plating” instructions above (steps 8 to 12). This step is particularly useful to know how well the cells survived in the microcentrifuge tube on ice during the transplantation procedure. Plating only three to five wells will be sufficient to determine viability, so a very small number of cells is needed for this step. SUPPORT PROTOCOL 2

PREPARATION OF COATED COVERSLIPS AND CULTURE FLASKS Coated coverslips and culture flasks are used to culture the neural progenitor cells.

Materials 100% ethanol 0.1 mg/ml poly-ornithine Round glass coverslips (Bellco, Fisher cat. no., NC9708845) or chamber slides 24-well tissue culture plates Tissue culture flasks 1. Wash coverslips with 100% ethanol and autoclave prior to use. Bellco round glass coverslips are optimal for the human neural progenitor cells plated in a 24-well plate. Chamber slides can also be used with appropriate volume modifications.

2. Under a laminar flow culture hood, place a coverslip in each well of a 24-well culture plate. 3. Coat each coverslip or each culture dish, plate, or flask with 0.1 mg/ml poly-ornithine and allow to sit in the culture hood at room temperature for at least 30 min. The volume of poly-ornithine needed will depend on the size of the well, plate, or flask, but use enough to completely cover the culture surface. For example, use 200 to 500 μl in each well of a 24-well plate or 1 to 10 ml in plates or flasks.

4. Remove poly-ornithine from each coverslip or culture surface and save for later use. Poly-ornithine can be reused three to five times.

5. Wash plates, flasks, or coverslips three times with sterile water and let dry in the hood overnight with the blower running. Do not expose coated plates and flasks to UV. If the UV must be used while the plates and flasks are drying in the hood, cover them with foil first. Plates and flasks can be stored up to 1 month at room temperature for later use. BASIC PROTOCOL 4

LENTIVIRAL INFECTION OF HUMAN NEURAL PROGENITOR CELLS FOR TRANSGENE OVEREXPRESSION Lentivirus can be used to infect human and rodent neural progenitor cells to stably integrate genes of interest. This procedure has been optimized for the human neural progenitor cells (Capowski et al., 2007) and will consistently produce ∼75% infection efficiency. The infection protocol described here is based on use of lentivirus with the phosphoglycerate kinase (PGK) promoter (Deglon et al., 2000; Dull et al., 1998; Naldini et al., 1996). Infections using other types of viruses or promoters will need to be optimized.

Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

CAUTION: All viruses should be handled with care and used in the appropriate biosafety level environment. Lentivirus can be neutralized with 70% ethanol, but check the environmental safety standards for clean-up and disposal when using other viruses.

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Materials Conditioned medium (CM; see Basic Protocol 3) Lentivirus Maintenance medium (see recipe) 70% ethanol 15- or 50-ml conical tubes 1.5-ml microcentrifuge tube Hemacytometer 24-well culture plate Additional reagents and equipment for complete dissociation of neural progenitor cells (Basic Protocol 3) Infect neural progenitor cells with lentivirus 1. Follow Basic Protocol 3, steps 1 to 7, to dissociate and count the cells. Retain the conditioned medium (CM). 2. Resuspend the cells in CM at 750 cells/μl in a 15-ml or 50-ml conical tube (depending on the number of wells to be used). A total of 300,000 cells will be needed for each well, plating a minimum of 10 wells for a total of 3 × 106 cells minimum. It is generally a good idea to calculate for an additional well to account for pipetting errors.

3. Add the appropriate volume of virus to the cell suspension in step 2 to infect the cells with the optimal viral titer. Lentiviral infection is based on the concentration of p24 coat protein. A standard infection is 100 ng p24 per million cells. This equates to 30 ng p24/well. The titer of the virus can vary widely in terms of infectious particles. The p24 coat protein assay identifies viral particles, but it does not show if the viral particles are infectious. The titer should be determined by quantitative PCR (Capowski et al., 2007). Furthermore, different cell types and viruses will need optimization to determine the appropriate infection titer.

Infect neural progenitor cells 4. Gently mix the virus/cell suspension and add 400 μl per well of a 24-well plate (for 300,000 cells/well). Incubate overnight. 5. The next day, add 600 μl fresh maintenance medium to each well, for a total of 1 ml per well. 6. Collect the small spheres 48 to 72 hr later in a 15-ml or 50-ml conical tube. All the cells that were infected with the same virus can be collected into the same tube. If multiple viruses were used, make sure to keep the cells with the different viruses separate. It is preferable to let spheres settle by gravity, but a gentle spin may be necessary. The rodent cells reform spheres very quickly, so they may need to be collected after 24 hr.

7. Resuspend in half fresh maintenance medium and half CM and seed into small flasks. Use a 12.5-cm2 flask when plating only 10 wells; larger flasks can be used when plating more wells, but keep the spheres denser than normal.

8. Split conservatively (1:1) into maintenance medium for the first 2 to 3 passages to allow the cells to recover. 9. Rinse tips and pipets that come into contact with virus in 70% ethanol prior to disposal. 10. Clean all work areas with 70% ethanol.

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ALTERNATE PROTOCOL 3

LENTIVIRAL INFECTION OF RODENT NEURAL PROGENITOR CELLS FOR TRANSGENE OVEREXPRESSION Rodent cells can be infected as dissociated cells within a flask rather than plating them into wells as described in Basic Protocol 4. Calculate the amount of virus needed to infect the number of cells, and add the virus directly to the flask. After the cells re-form spheres (in ∼48 to 72 hr), remove all the medium and replace with fresh maintenance medium.

BASIC PROTOCOL 5

CRYOPRESERVATION AND SUBSEQUENT THAWING OF NEURAL PROGENITOR CELLS FOR LONG-TERM STORAGE Human and rodent neural progenitor cells can be stored indefinitely in frozen stocks in liquid nitrogen without impacting their growth properties upon thawing and re-culturing. This is useful for generating cell banks for large-scale experiments.

Materials Human or rodent neurospheres (Basic Protocol 1) Maintenance or starting medium (see recipe) Cell Freezing Medium (Sigma, cat no. C6295) Liquid N2 Base medium (see recipe) Starting medium (see recipe) Cryovials Isopropanol freezing chamber 15-ml conical tube Liquid nitrogen storage tank Additional reagents and equipment for complete dissociation of neural progenitor cells (Basic Protocol 3) and counting viable cells by trypan blue exclusion (UNIT 1C.3) Freeze cells 1. Follow steps 1 to 7 in Basic Protocol 3, substituting trypsin for Accutase in step 2 and maintenance or starting medium for plating medium in step 5. 2. Using a hemacytometer, count an aliquot of cells and assess viability using trypan blue (UNIT 1C.3). Follow a similar dilution as described in step 8 of Basic Protocol 1.

3. Microcentrifuge 3 min at 3000 × g to pellet the cells. The cells can be spun faster, but avoid that if possible.

4. Remove and discard the medium and gently resuspend in cell freezing medium. Add the appropriate volume of freezing medium for a final cell concentration of 5 million cells/ml. Sigma Cell Freezing medium is optimal for the human and rodent progenitor cells. The freezing medium is only good for 3 to 5 days after opening.

5. Transfer 1 ml of the cell suspension to each cryovial and incubate 10 min on ice. 6. Transfer the cryovials to a room temperature isopropanol freezing chamber and place in a −80◦ C freezer overnight. Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

7. Transfer each cryovial to a liquid nitrogen storage unit after ∼24 hr.

Thaw cells 8. Add 10 ml of base medium to a 15-ml conical tube. Use one conical tube for each cryovial.

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9. Place the cryovial in a 37◦ C water bath very briefly, just until the cells have started to thaw. Thaw only a few tubes at once.

10. Add the contents of the cryovial into the conical tube, allow the cells to settle, and remove the medium. To pellet the cells, a brief (and gentle) microcentrifugation (e.g., 3 min at 3000 × g) may be necessary.

11. Wash the cells with 10 ml base medium a second time. Because the freezing medium contains DMSO, it is important to thoroughly wash the cells.

12. Add 10 ml of starting medium to each conical tube and gently resuspend the cells. Use starting medium for both human and rodent cells. LIF should be added if the human cells are at the appropriate passage number (e.g., >passage 10).

13. Transfer the cells to the appropriate-sized flask at a concentration of 200,000 cells/ml. Rodent and human cells should reform spheres within 2 to 3 days. Depending on the health and size of the spheres, cells can be transferred to maintenance medium 1 to 2 weeks after the thaw.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Base medium 70% (v/v) Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 4500 mg/liter glucose plus L-glutamine and NaHCO3 30% (v/v) Ham’s F-12 nutrient solution Antibiotic/antimycotic solution (penicillin/streptomycin/amphotericin B, PSA) to 1× final All base medium components can be purchased from various vendors and stored unopened or in frozen aliquots according to manufacturers’ instructions. Once the base medium is made, it can be stored at 4◦ C for up to a month. Stemline (Sigma, cat. no. S3194), with added PSA, can be used as an alternative base medium. However, note that Stemline already contains supplements, so if using it as the base for the other medium described below, it is not necessary to add additional B27 or N2 supplements but they can be added without detriment. Stemline has been optimized for human progenitor cell cultures, although it is suitable for cultures derived from other species.

Maintenance medium Base medium (see recipe) supplemented with: 1% (v/v) N2 supplement (Invitrogen) 20 ng/ml epidermal growth factor (EGF) 10 ng/ml leukemia inhibitory factor (LIF; Millipore; only add for human cultures older than passage 10) Maintenance medium can be stored at 4◦ C for 2 weeks. EGF can be purchased from various vendors.

Plating medium Base medium (see recipe) supplemented with: 2% (v/v) B27 supplement (Invitrogen) 1% (v/v) fetal bovine serum (FBS; optional) Plating medium can be stored at 4◦ C for 2 weeks.

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Starting medium Base medium (see recipe) supplemented with: 2% (v/v) B27 supplement (Invitrogen) 20 ng/ml epidermal growth factor (EGF) 20 ng/ml fibroblast growth factor 2 (FGF-2) 5 μg/ml heparin Starting medium can be stored at 4◦ C for 2 weeks. EGF, FGF, and heparin can be purchased from various vendors. Heparin is added to any medium containing FGF-2 because, among other things, it stabilizes the FGF-2 and promotes faster growth (Caldwell et al., 2004).

Transplant medium 50% (v/v) Leibowitz (L-15) medium 50% (v/v) 0.6% (w/v) glucose in PBS without Ca or Mg (Invitrogen, cat. no. 14190-250) 2% (v/v) B27 supplement (Invitrogen) The transplant medium is best made up fresh. The Leibowitz medium can be purchased from various vendors and stored according to the manufacturers’ instructions. Opened bottles can be stored at 4◦ C until the expiration date.

COMMENTARY Background Information

Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

There are various applications for neural progenitor cells. These include studying migration, proliferation, and differentiation of particular neural cell populations in vitro and transplantation into rodent central nervous system to determine cellular characteristics in vivo. They can also be used to develop novel transplant therapies for diseases of the brain and spinal cord. Human and rodent progenitor cell cultures provide an essentially limitless source of neural material because of their rapid rate of expansion. Neurosphere cultures are a convenient way to propagate the cells due to the large number of spheres that can be cultured in each flask compared to plated cells. However, they also have an increased complexity due to spontaneous differentiation within the neurospheres. Monolayer cultures are convenient and easy to grow, but they may show different characteristics than neurosphere cultures. One could argue that the three-dimensional environment of the neurosphere mimics the in vivo situation when compared to the artificial nature of twodimensional culture systems. However, once a cell is removed from its in vivo environment, everything becomes an artifact. Therefore, each culture system should be taken at face value. Progenitor cells derived from different regions of the central nervous system and at different stages of development have different properties, and may respond better

to one growth condition compared to another. Therefore, the optimal growth method and conditions will need to be determined by the end user based on experimental needs.

Troubleshooting Table 2D.2.1 provides troubleshooting information for neural progenitor cell protocols. Mouse embryonic cells in particular exhibit a tendency toward chromosomal instability in culture (Todaro and Green, 1963). Therefore, it is recommended the mouse cultures be discarded after 5 to 6 passages.

Anticipated Results When starting with eight mouse or rat embryos, 4 to 8 million cells will be collected from the developing striatum. These can be seeded at 200,000 cells/ml until the first passage, at which time they are seeded at 100,000 cells/ml. Due to the rapid expansion, the number of flasks/plates can double every 4 to 7 days. For the human cells, it is best to keep the cells dense. For example, if a cryotube containing 5 million human progenitor cells is thawed, seed into a 25-cm2 flask until they reform spheres; some cell death is expected (∼10% to 20%). Generally, a dense 25-cm2 flask will have 3 to 5 million cells; a dense 75-cm2 flask will have 10 to 15 million cells; and a dense 175-cm2 flask will have 20 to 25 million cells.

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Table 2D.2.1 Troubleshooting Guide to Neural Progenitor Cell Protocols

Problem

Possible cause

Solution

Neurospheres stop growing or begin to stick to the flask

Too old

Discard cells and thaw out younger neurospheres. Human cells will eventually senesce, so this is not unexpected as the cultures reach 50 or more passages (Wright et al., 2006).

Cells do not re-form spheres after chop

Chopped too small

Allow neurospheres to become 500-700 mm prior to chopping, and confirm that chopper is set to 200 mm.

Cells die after plating/dissociation

Enzyme left on cells too long and/or cells triturated too harshly

Thaw enzyme just prior to use and do not leave on the cells longer than 10 min. If the cells have not completely dissociated after ∼100 passes through a pipet tip, stop triturating. More damage will be done to the cells by continual mechanical dissociation than by having a few small clumps remaining.

The percent of neurons, astrocytes, and oligodendrocytes generated following terminal differentiation will depend on the region from which the cells were derived and the passage number. Cells will generally continue to follow an intrinsic developmental pattern of generating more neurons in early passages and then generating more astrocytes at later passages. This may reflect the fact that within growing neurospheres or plated cultures there are only a few “true” self-renewing stem cells surrounded by many committed progenitors. New methods to isolate and grow “true” stem cells should be developed, which may be dependent upon learning more about stem cell niches and self-renewal (Alvarez-Buylla and Lim, 2004). Infection with lentivirus will maintain stable integration for many weeks and passages. We have found lentiviral-induced growth factor overexpression from the human cortical progenitor cells to persist for at least 3 months in rats and monkeys (Behrstock et al., 2006). If properly stored in liquid nitrogen, frozen cell stocks are viable indefinitely.

Lentiviral infection takes 30 to 45 min/ sample. Again, when comfortable, multiple samples and infections can be done at the same time.

Time Considerations

Chopping takes ∼15 min/flask, not including the initial chopper setup or UV sterilization prior to starting or between samples. Enzymatic dissociation takes ∼30 min/ sample, including incubation times. When comfortable, multiple samples can be dissociated at the same time, thus decreasing total time.

Literature Cited Alvarez-Buylla, A. and Lim, D.A. 2004. For the long run: Maintaining germinal niches in the adult brain. Neuron 41:683-686. Behrstock, S., Ebert, A., McHugh, J., Vosberg, S., Moore, J., Schneider, B., Capowski, E., Hei, D., Kordower, J., Aebischer, P., and Svendsen, C.N. 2006. Human neural progenitors deliver glial cell line-derived neurotrophic factor to parkinsonian rodents and aged primates. Gene Ther. 13:379-388. Caldwell, M.A., Garcion, E., ter Borg, M.G., He, X., and Svendsen, C.N. 2004. Heparin stabilizes FGF-2 and modulates striatal precursor cell behavior in response to EGF. Exp. Neurol. 188:408-420. Capowski, E.E., Schneider, B.L., Ebert, A.D., Seehus, C.R., Szulc, J., Zufferey, R., Aebischer, P., and Svendsen, C.N. 2007. Lentiviral vectormediated genetic modification of human neural progenitor cells for ex vivo gene therapy. J. Neurosci. Methods 163:338-349. Chandran, S., Compston, A., Jauniaux, E., Gilson, J., Blakemore, W., and Svendsen, C. 2004. Differential generation of oligodendrocytes from human and rodent embryonic spinal cord neural precursors. Glia 47:314-324. Deglon, N., Tseng, J.L., Bensadoun, J.C., Zurn, A.D., Arsenijevic, Y., Pereira, d.A., Zufferey, R., Trono, D., and Aebischer, P. 2000. Selfinactivating lentiviral vectors with enhanced transgene expression as potential gene transfer system in Parkinson’s disease. Hum. Gene Ther. 11:179-190.

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Dull, T., Zufferey, R., Kelly, M., Mandel, R.J., Nguyen, M., Trono, D., and Naldini, L. 1998. A third-generation lentivirus vector with a conditional packaging system. J. Virol. 72:84638471. Hitoshi, S., Tropepe, V., Ekker, M., and van der Kooy, D. 2002. Neural stem cell lineages are regionally specified, but not committed, within distinct compartments of the developing brain. Development 129:233-244.

neurons. Proc. Natl. Acad. Sci. U.S.A. 90:36023606. Reynolds, B.A. and Weiss, S. 1992. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707-1710. Reynolds, B.A. and Weiss, S. 1996. Clonal and population analyses demonstrate that an EGFresponsive mammalian embryonic CNS precursor is a stem cell. Dev. Biol. 175:1-13.

Johe, K.K., Hazel, T.G., Muller, T., DugichDjordjevic, M.M., and McKay, R.D. 1996. Single factors direct the differentiation of stem cells from the fetal and adult central nervous system. Genes Dev. 10:3129-3140.

Svendsen, C.N., Fawcett, J.W., Bentlage, C., and Dunnett, S.B. 1995. Increased survival of rat EGF-generated CNS precursor cells using B27 supplemented medium. Exp. Brain Res. 102:407-414.

Kim, H.T., Kim, I.S., Lee, I.S., Lee, J.P., Snyder, E.Y., and Park, K.I. 2006. Human neurospheres derived from the fetal central nervous system are regionally and temporally specified but are not committed. Exp. Neurol. 199:222-235.

Svendsen, C.N., Skepper, J., Rosser, A.E., ter Borg, M.G., Tyres, P., and Ryken, T. 1997. Restricted growth potential of rat neural precursors as compared to mouse. Brain Res. Dev. Brain Res. 99:253-258.

Laywell, E.D., Rakic, P., Kukekov, V.G., Holland, E.C., and Steindler, D.A. 2000. Identification of a multipotent astrocytic stem cell in the immature and adult mouse brain. Proc. Natl. Acad. Sci. U.S.A. 97:13883-13888.

Svendsen, C.N., terBorg, M.G., Armstrong, R.J., Rosser, A.E., Chandran, S., Ostenfeld, T., and Caldwell, M.A. 1998. A new method for the rapid and long term growth of human neural precursor cells. J. Neurosci. Methods 85:141152.

Morshead, C.M., Benveniste, P., Iscove, N.N., and van der Kooy, D. 2002. Hematopoietic competence is a rare property of neural stem cells that may depend on genetic and epigenetic alterations. Nat. Med. 8:268-273. Naldini, L., Blomer, U., Gallay, P., Ory, D., Mulligan, R., Gage, F.H., Verma, I.M., and Trono, D. 1996. In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263-267. Ostenfeld, T., Joly, E., Tai, Y.T., Peters, A., Caldwell, M., Jauniaux, E., and Svendsen, C.N. 2002. Regional specification of rodent and human neurospheres. Brain Res. Dev. Brain Res. 134:43-55. Ray, J. and Gage, F.H. 1994. Spinal cord neuroblasts proliferate in response to basic fibroblast growth factor. J. Neurosci. 14:3548-3564. Ray, J. and Gage, F.H. 2006. Differential properties of adult rat and mouse brain-derived neural stem/progenitor cells. Mol. Cell Neurosci. 31:560-573. Ray, J., Peterson, D.A., Schinstine, M., and Gage, F.H. 1993. Proliferation, differentiation, and long-term culture of primary hippocampal

Todaro, G.J. and Green, H. 1963. Quantitative studies of the growth of mouse embryo cells in culture and their development into established lines. J. Cell Biol. 17:299-313. Vescovi, A.L., Parati, E.A., Gritti, A., Poulin, P., Ferrario, M., Wanke, E., Frolichsthal-Schoeller, P., Cova, L., Arcellana-Panlilio, M., Colombo, A., and Galli, R. 1999. Isolation and cloning of multipotential stem cells from the embryonic human CNS and establishment of transplantable human neural stem cell lines by epigenetic stimulation. Exp. Neurol. 156:71-83. Watanabe, K., Nakamura, M., Iwanami, A., Fujita, Y., Kanemura, Y., Toyama, Y., and Okano, H. 2004. Comparison between fetal spinal-cordand forebrain-derived neural stem/progenitor cells as a source of transplantation for spinal cord injury. Dev. Neurosci. 26:275-287. Wright, L.S., Prowse, K.R., Wallace, K., Linskens, M.H., and Svendsen, C.N. 2006. Human progenitor cells isolated from the developing cortex undergo decreased neurogenesis and eventual senescence following expansion in vitro. Exp. Cell Res. 312:2107-2120.

Isolating, Expanding, and Infecting Human and Rodent Fetal Neuroprogenitor Cells

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Long-Term Multilayer Adherent Network (MAN) Expansion, Maintenance, and Characterization, Chemical and Genetic Manipulation, and Transplantation of Human Fetal Forebrain Neural Stem Cells

UNIT 2D.3

Dustin R. Wakeman,1, 2 Martin R. Hofmann,2 D. Eugene Redmond, Jr.,3 Yang D. Teng,4, 5 and Evan Y. Snyder1, 2 1

University of California at San Diego, La Jolla, California The Burnham Institute for Medical Research, La Jolla, California 3 Yale University School of Medicine, New Haven, Connecticut 4 Harvard Medical School, Brigham & Women’s Hospital and Spaulding Rehabilitation Hospital, Boston, Massachusetts 5 Veterans Affairs Boston Healthcare System, Boston, Massachusetts 2

ABSTRACT Human neural stem/precursor cells (hNSC/hNPC) have been targeted for application in a variety of research models and as prospective candidates for cell-based therapeutic modalities in central nervous system (CNS) disorders. To this end, the successful derivation, expansion, and sustained maintenance of undifferentiated hNSC/hNPC in vitro, as artificial expandable neurogenic micro-niches, promises a diversity of applications as well as future potential for a variety of experimental paradigms modeling early human neurogenesis, neuronal migration, and neurogenetic disorders, and could also serve as a platform for small-molecule drug screening in the CNS. Furthermore, hNPC transplants provide an alternative substrate for cellular regeneration and restoration of damaged tissue in neurodegenerative disorders such as Parkinson’s disease and Alzheimer’s disease. Human somatic neural stem/progenitor cells (NSC/NPC) have been derived from a variety of cadaveric sources and proven engraftable in a cytoarchitecturally appropriate manner into the developing and adult rodent and monkey brain while maintaining both functional and migratory capabilities in pathological models of disease. In the following unit, we describe a new procedure that we have successfully employed to maintain operationally defined human somatic NSC/NPC from developing fetal, pre-term postnatal, and adult cadaveric forebrain. Specifically, we outline the detailed methodology for in vitro expansion, long-term maintenance, manipulation, and transplantation of these C 2009 by John multipotent precursors. Curr. Protoc. Stem Cell Biol. 9:2D.3.1-2D.3.77.  Wiley & Sons, Inc. Keywords: human neural stem/progenitor cell r NPC r NSC r culture r fetal/adult forebrain r subventricular zone r neurogenesis r niche r multilayer adherent network r MAN assay r protocols r manipulation techniques r characterization r in vitro r derivation r expansion r maintenance r SPIO r Feridex r lentivirus r BrdU r labeling

INTRODUCTION A number of techniques have been devised to attempt to identify and isolate rodent and human neural stem/precursor cells (NSCs/NPCs). Some have relied on the aggregation of cells in suspension cultures—termed “neurospheres” and giving rise to the “neurosphereforming assay” (NSA; Reynolds and Weiss, 1992; Reynolds et al., 1992; Rietze and Reynolds, 2006)—for artificially expanding nonclonal NSC/NPC populations in vitro Current Protocols in Stem Cell Biology 2D.3.1-2D.3.77 Published online May 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d03s9 C 2009 John Wiley & Sons, Inc. Copyright 

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(Singec et al., 2006) in serum-free medium. However, other techniques have been employed prior to (Ryder et al., 1990; Redies et al., 1991; Renfranz et al., 1991; Snyder et al., 1992) and since (Flax et al., 1998; Shihabuddin et al., 1996; Rubio et al., 2000) popularization of the NSA which, in fact, have been found to have beneficial properties compared to the NSA. It is these techniques that our group has long employed to great advantage and success—particularly when interested in using NSCs/NPCs for transplantation, genetic manipulation, rigorous clonal analyses, and developmental studies—and which will be described in this unit. Human embryonic, fetal, newborn, and adult cadaveric CNS precursors have been shown to thrive when derived and maintained as two-dimensional (2-D) adherent cultures. This technique offers many growth and culture advantages over the NSA and, in fact, has come to supplant the NSA in many neurobiological laboratories. Over the past two decades, numerous techniques have been described for the derivation and expansion of suspension of human neural precursors either in suspension or as adherent monolayers (Ray et al., 1995; Svendsen et al., 1999; Wu et al., 2002; Walsh et al., 2005; Rajan and Snyder, 2006; Ray and Gage, 2006; Pollard et al., 2006a,b), utilizing an assortment of growth factors (Buc-Caron, 1995; Chalmers-Redman et al., 1997; Moyer et al., 1997; Sah et al.,1997; Svendsen et al., 1998, 1999; Carpenter et al., 1999; Kukekov et al., 1999; Vescovi et al., 1999a,b; Roy et al., 2000; Uchida et al., 2000; Villa et al., 2000; Piper et al., 2000, 2001; Arsenijevic et al., 2001a,b; Keyoung et al., 2001; Palmer et al., 2001; Cai et al., 2002; Laywell et al., 2002; Nunes et al., 2003; Schwartz et al., 2003; Zhang et al., 2005; Conti et al., 2005; Li et al., 2005; Pollard et al., 2006a,b; Yin et al., 2006; Ray, 2008). In this unit, we outline methodology for the expansion, long-term maintenance, manipulation, and transplantation of human fetal (10- to 25-week) neural precursor cells (hNPC). Specifically, we describe a new method for long-term expansion of karyotypically stable hNPC, termed the Multilayer Adherent Network (MAN), to generate largescale self-renewing multipotent hNPC populations, amenable to in vitro manipulation and transplantation in vivo. We describe in detail the methods we have successfully utilized to prepare and transplant hNPC into the neonatal mouse and adult nonhuman primate. In addition, we provide basic procedures for characterization of undifferentiated and differentiated hNPC, as well as the processing of engrafted brains. Furthermore, we illustrate techniques for the efficient labeling of hNPC, including lentivirus infection and noninvasive superparamagnetic iron oxide (SPIO) particle transfection. For simplicity’s sake, we will refrain from the operational NSC debate and simply refer to both neural stem and progenitor cells as NPCs from here forward. The protocols in order of presentation are: Basic Protocol 1: Establishing and maintaining multilayer adherent network (MAN) cultures; Support Protocol 1: Derivation of human fetal neural stem/precursor cells; Alternate Protocol 1: Feeding and dissociation of lightly adherent aggregate cultures; Alternate Protocol 2: Growing hNPC in MAN membrane system (MMS); Support Protocol 2: Cryopreservation of hNPC; Support Protocol 3: Thawing cryopreserved hNPC; Support Protocol 4: Preservation of conditioned medium; Long-Term MAN Growth and Characterization of NPCs

Alternate Protocol 3: Replating dissociated hNSC on extracellular matrix (ECM) as adherent two-dimensional monolayer cultures; Support Protocol 5: Preparation of extracellular matrix (ECM) substrates;

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Basic Protocol 2: Establishing clonal hNPC subpopulations; Basic Protocol 3: Labeling hNPC with BrdU; Basic Protocol 4: Lentiviral infection of hNPC; Alternate Protocol 4: Lentiviral infection of multilayer adherent network (MAN); Basic Protocol 5: Labeling hNPC with super-paramagnetic iron oxide (SPIO); Support Protocol 6: Perls Prussian blue staining (for hemosiderin); Basic Protocol 6: Preparing hNPC for transplantation; Basic Protocol 7: Loading and injection of hNPC for transplantation into St. Kitts African Green Monkey; Basic Protocol 8: Intraventricular injection of hNPC into neonatal mice; Basic Protocol 9: Processing engrafted mouse brains; Basic Protocol 10: Characterizing hNPC. NOTE: The following procedures are performed aseptically in a sterile, Biosafety Level 2 hood. NOTE: A standard pathogen testing program for hepatitis B and C, HTLV-1/2, syphilis RPR, HIV-1/2, cytomegalovirus, Hantaviruses (Hantaan, Seoul, Sin Nombre), West Nile virus, Trypanosoma cruzi, and mycoplasma should be carried out throughout the entire natural history of the NPC culture to ensure proper safety. We recommend the human IMPACT Profile pathogen test in conjunction with the IMPACT Profile VIII: Comprehensive Murine Panel from the University of Missouri Research Animal Diagnostic Laboratory (RADIL) to monitor hNPC populations throughout long-term expansion. NOTE: Periodic cytogenetic testing for acquisition of gross chromosomal alteration in vitro is also recommended to confirm a normal human karyotype complement.

STRATEGIC PLANNING Growth Factor Signaling Long-term expansion and maintenance of self-renewing NPC in serum-free media (Reynolds et al., 1992; Reynolds and Weiss, 1992; Svendsen et al., 1996; Rosser et al., 1997) requires mitogenic support from either epidermal growth factor (EGF) or basic fibroblast growth factor (bFGF) to activate mitogen-activated-protein-kinase (MAPK) signaling and support hNPC division (Gensburger et al., 1987; Walicke, 1988; Kornblum et al., 1990; Murphy et al., 1990; Drago et al., 1991a,b; Ray et al., 1993; Vescovi et al., 1993a,b; Bartlett et al., 1994; Kitchens et al., 1994; Ray and Gage, 1994; Ghosh and Greenberg, 1995; Kilpatrick and Bartlett, 1993, 1995; Kilpatrick et al., 1995; Palmer et al., 1995; Vicario-Abejon et al., 1995; Gritti et al., 1996; Kuhn et al., 1997; Qian et al., 1997; Shihabuddin et al., 1997; Caldwell and Svendsen, 1998; Ciccolini and Svendsen, 1998; Gritti et al., 1999; Palmer et al., 1999; Arsenijevic et al., 2001a,b; Caldwell et al., 2001; Temple, 2001; Ostenfeld and Svendsen, 2004; Tarasenko et al., 2004; Kelly et al., 2005; Ray and Gage, 2006). In addition, the neurotrophic leukemia inhibitory factor (LIF) has been shown to enhance telomerase expression, improve viability, and extend the time until terminal senescence of hNPC when used in combination with bFGF and/or EGF (Galli et al., 2000; Molne et al., 2000; Shimazaki et al., 2001; Wright et al., 2003; Bonaguidi et al., 2005; Gregg and Weiss, 2005). Although LIF signaling appears to induce gliogenesis in rodent NPC, in our experience, LIF not only enhances survival and

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doubling time of human NPC but is absolutely essential for the sustained maintenance of symmetric cell divisions in long-term multilayer adherent network cultures. Direct comparisons of NPC derived from different species or by alternate techniques have shown that NPC characteristics are drastically altered by their environmental inputs and retain these intrinsic cellular properties in direct relation to how they are manipulated in vitro (Ray and Gage, 2006). We have empirically determined the specific regimen of growth factors that best supports growth of human fetal forebrain NPC. As a result, we have adopted a strategy for sustained proliferative expansion of karyotypically normal undifferentiated hNPC in basal growth medium consisting of bFGF and LIF (without EGF).

Media Formulations Although traditional serum-free rodent NPC culture has generally utilized DMEM/F12 supplemented with N2, we have adjusted the recipe to accommodate hNPC by utilizing Neurobasal medium (Invitrogen) with B-27 supplement (without vitamin A) to support long-term proliferation of hNPC in vitro (Brewer et al., 1993; Brewer, 1995, 1997; Svendsen et al., 1995; Brewer and Price, 1996; Brewer and Torricelli, 2007). In addition, heparin is added to stabilize the binding of the bFGF heparin-sulfate proteoglycan to its FGFR-1 receptor (Balaci et al., 1994; Caldwell et al., 2004), potentiating cell-cell attachments that favor adherent monolayer hNPC growth (Richard et al., 1995, 2000). On the day of use, prepare fresh hNPC growth medium plus 20 ng/ml bFGF plus 10 ng/ml LIF (see Reagents and Solutions). Growth factors are added fresh on the day of use due to their relative instability (Kanemura et al., 2005). Contamination is possible and thus Normocin (InvivoGen) is supplemented regularly (48-hr half-life) as an antipathogenic agent (replaces penicillin/streptomycin/amphotericin B to deter mycoplasma, Grampositive and -negative bacteria, and fungal contamination). Normocin and any other antibiotics employed may be gradually withdrawn from the culture after an adequate period of time as desired. Due to the relatively large amount of time and resources involved in hNPC culture, we highly recommend the use of pathogen-control agents. Normocin has remained the most gentle yet potent and comprehensive single treatment application we have tested thus far. LONG-TERM EXPANSION AND MAINTENANCE OF hNPC Throughout the expansion process, cryopreservation and functional testing of hNPC lines is necessary for the continued long-term maintenance of healthy proliferative progenitors. Cultures are monitored superficially under the light microscope for morphological aberrations that may occur in artificial culture. Once sufficient cell numbers have been established, a more intensive battery of screens for in vitro and in vivo multipotency should be employed, particularly when hNPC reach high passage number or whenever a new vial of early passage progenitors are thawed from cryopreservation for mass expansion, to ensure hNPC cultures do not change phenotypically or become lineage restricted with time. To test functionality, several vials are reconstituted to assess the overall freeze/thaw success, cell viability, and sustained multipotency. Throughout culture, the genetic stability of hNPC should be confirmed periodically through spectral karyotyping, microarray fingerprinting, and transcriptome and proteomic analysis to demonstrate a normal chromosomal complement and sustained expression profile of all classical stemness-associated genes (Cai et al., 2006; Chang et al., 2006; Luo et al., 2006a,b; Maurer and Kuscinsky, 2006; Shin and Rao, 2006; Anisimov et al., 2007; Shin et al., 2007). In an effort to reduce time and costly resources, hNPC lines should be regularly tested for these attributes before proceeding with any large animal transplantation studies. Long-Term MAN Growth and Characterization of NPCs

NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator, unless otherwise noted.

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Table 2D.3.1 Plating Volumes for Different Culture Vessels

Area (cm2 /well)

Vessel

Volume

Petri dishes 20 mm

3

1 ml

25 mm

8

2.5 ml

60 mm

25

6 ml

100 mm

78.5

18 ml

6 well

9.6

3.5 ml

12 well

3.8

2 ml

24 well

2

1 ml

48 well

0.75

500 μl

96 well

0.32

250 μl

1 well

9.4

3 ml

2 well

4.2

2 ml

4 well

1.8

1 ml

8 well

0.8

250 μl

25

6-8 ml

75

16-20 ml

225

40-50 ml

Multiwell plates

Slides

Flask 25-cm2 2

75-cm

2

225-cm

NOTE: All reagents and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: Numerous different types and sizes of tissue culture vessels are described in this unit; the plating volumes for common tissue culture petri dishes, multiwell plates, slides, and flasks are listed in Table 2D.3.1.

ESTABLISHING AND MAINTAINING MULTILAYER ADHERENT NETWORK (MAN) CULTURES Traditionally, we have thawed and grown hNSC as small, slightly adherent aggregates for the first 2 to 3 weeks of culture post-thaw. More recently, however, we have developed a new method for expansion of newly thawed or freshly dissociated undifferentiated hNPC on noncoated flasks free of extracellular matrix (ECM). Establishment of these high-density multilayer adherent networks (MAN) is founded on the basic theory of aggregate formation, but is adapted into a novel adherent system that offers many growth advantages for both the progenitor population and the researcher. As a whole, the MAN assay relies on a combination of the inherent hNPC property of forming fusion aggregates at high density, coupled with the intrinsic behavior of resting hNPC aggregates to attach and migrate over time. The end result is a highly dynamic, proliferative population of undifferentiated hNPC displaying a variety of advantageous growth parameters. In general, we find that mature MAN hNPC cultures proliferate and expand at an elevated doubling rate (3 to 5 days) compared to their neurosphere counterparts (4 to 7 days; Kanemura et al., 2002, 2005; Mori et al., 2006). In addition, feeding MAN cultures fresh medium can easily be achieved by simply tilting the flask, aspirating or collecting

BASIC PROTOCOL 1

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CM, and refilling the flask with new medium. This fast and easy process allows the researcher to replace all or portions of the medium as often as necessary without the harsh mechanical stresses involved in centrifugation. The key to transitioning traditional aggregate cultures into MAN cultures is the overall density of the hNPC initially plated. Simply stated, the greater the density of hNPC initially plated, the larger the aggregate units, the more quickly they attach, and, thus, the more quickly subsequent mature multilayer adherent networks are established. It should be noted that replating hNPC at densities greater than 4 × 106 cells per 25-cm2 flask will result in overcrowding and subsequent formation of large spheroid cellular masses, negating the entire premise for the initial dissociation. For the most part, highdensity passaging is only recommended for preparing small cellular clusters prior to cryopreservation, or to quickly establish mature MAN cultures for short-term study. A brief history of the early stages of MAN formation is: a. 0 to 24 hr: Cells equilibrate and settle to bottom of flask following an even distribution pattern. b. 24 to 48 hr: Cells begin to lightly attach and spread (as evidenced by small microspikes and several small projections; Fig. 2D.3.1A,B). c. 48 to 72 hr: Aggregated cell clusters continue to spread, elongate, and begin to proliferate and extend into adjacent neighboring clusters, becoming adherent three-dimensional clusters, creating the first evidence of an interlinked network (Fig. 2D.3.1C). d. 72 to 96 hr: Cell clusters continue to migrate into each other at the periphery and become anchored strongly enough to change medium. These cultures consist mainly of slightly adherent clusters and a small proportion of nonadherent floating aggregates. The cultures can be carefully removed from the incubator to change medium without disrupting the newly formed MAN (Fig. 2D.3.1D-F).

Materials Human NPC (Support Protocol 1): frozen (Support Protocol 2) and freshly thawed (Support Protocol 3) or freshly dissociated as described in Support Protocol 1 25% (v/v) conditioned medium (CM; Support Protocol 4)/75% (v/v) NB-B-27 complete medium (see recipe) containing 40 ng/ml bFGF and 10 ng/ml LIF (bFGF and LIF concentrations based on total volume of medium) NB-B-27 complete medium (see recipe) Leukemia inhibitor factor (LIF; Millipore, cat. no. LIF1010) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) Normocin (InvivoGEN, cat. no. ant-nr-1) Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) Dulbecco’s PBS without Ca2+ or Mg2+ (CMF-DPBS; Mediatech, cat. no. 21-031-CM) Accutase (Millipore, cat. no. SCR005) or Cell Dissociation Buffer (Invitrogen, cat. no. 13150-016) Conditioned medium (CM; Support Protocol 4)

Long-Term MAN Growth and Characterization of NPCs

15-ml conical tubes 25-cm2 and 75-cm2 tissue culture flasks Battery-powered pipetting aid (e.g., Drummond Pipet-Aid XP) 1000-μl extended-length pipet tip and 1000-μl automatic pipettor Centrifuge Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3)

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Figure 2D.3.1 Establishment of multilayer adherent network (MAN). (A) 24 to 48 hr after plating, hNSC (HFB2050) readily form evenly spaced, proliferative aggregated cell clusters. Small hNPC clusters initially attach to the culturing surface and sample the local microenvironment with meandering growth-cone like protrusions (B), and eventually flatten and spread out (C). Taking advantage of higher plating densities, the MAN culturing technique creates optimal spacing between colonies, allowing each aggregate cluster close access to neighboring signaling molecules. (D-F) After 72 hr, hNSC aggregates are lightly attached to the surface and begin to actively proliferate. Over the next 3 to 4 weeks, hNSC rapidly expand and form extensive honeycomb-shaped, mature multilayer adherent networks (G-I).

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Figure 2D.3.2 Human neural precursor cell basic culture schedule. Human NPC are grown as either lightly adherent aggregates or as multilayer adherent networks in 25-cm2 flasks. Conditioned medium (CM) is gradually reduced from cultures as they progress and can be collected after MAN cultures reach ∼75% confluence. Aggregate cultures are dissociated once a week or as growth parameters dictate, whereas MAN cultures can be cultured for up to 1 month before passaging.

Establish MAN cultures 1. To establish MAN cultures from freshly thawed cells or freshly dissociated cells, resuspend hNPC 2:1 (i.e., at 2–3 × 106 cells/flask) in 25% (v/v) CM/75% (v/v) NB-B-27 complete medium (containing 40 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin) in a 15-ml conical tube. Transfer hNPC to an uncoated 25-cm2 flask (Fig. 2D.3.2). Ratios such as 2:1 refer to the surface area used—i.e., if starting with one 25-cm2 flask, when expanding cells, one would use a 1:2 split, meaning that one should start with one 25-cm2 flask and resuspend the dissociated cells into two 25-cm2 flasks—increasing the surface area from 25 to 50 or 1:2. However, if referring to establishment of a culture with frozen cells, the ratio is 2:1, i.e., the number of frozen cells that were originally in two 25-cm2 flasks would need to be thawed into one 25-cm2 flask. Similarly, when dealing with freshly dissociated cells, the ratio is 2:1. In this step, 2 million fresh cells or 3 million frozen cells are diluted into 8 ml media into one 25-cm2 flask. Plating a higher density of hNPC leads to the quicker (24- to 72-hr) formation of small (2–3 × 106 cells) to medium size (3–4 × 106 cells) clusters, respectively, initiating close cell-cell contacts critical for enhanced paracrine and autocrine support. This means that if you plate 2–3 × 106 cells (dissociated) into one 25-cm2 flask, it will give you small clusters within 24 to 72 hr, whereas 3–4 × 106 will give medium-size clusters in this same period of time. Interestingly, we have found that leukemia inhibitory factor (LIF) is absolutely necessary and essential for the long-term maintenance of MAN cultures. Removal of LIF from the basal growth medium results in the rapid breakdown of elongated projections into ropelike, flexible, spindly, nonadherent protrusions that eventually disappear, ultimately resulting in the loss of proliferation capacity, increased senescence, and eventual cellular crisis.

2. Place the flask into a humidified 5% CO2 incubator at 37◦ C and shake horizontally in both planes to evenly disperse cells throughout the flask without sloshing medium into the neck of the flask. Long-Term MAN Growth and Characterization of NPCs

It is imperative that small (4- to 16-cell) to medium size (16- to 64-cell) clusters (from thawed sample; Support Protocol 3) or single cells (from dissociation) are dispersed uniformly onto the surface to avoid clumping and uneven coating of the flask.

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3. Once the cells have been fully distributed, allow the flask to incubate and equilibrate for 3 days without moving the flask for any reason from its original resting position. It is equally important that the incubator remain motionless and not be bumped or shaken in any way, or else the even hNPC coating will be disrupted and the organization of the MAN system will become disordered. During the 72-hr hands-free period, the individual evenly spaced cellular clusters settle to the bottom of the flask, lightly attach, and migrate out over the surface, proliferating into each other, creating an interlinked lattice of three-dimensional adherent clusters displaying elongated processes that extend and connect each cellular island into a global multilayer adherent network (MAN). Perhaps the most important aspects of successful MAN culture setup are the initial plating conditions coupled with diligent patience and a steady hand during the initial week after hNPC thaw. Throughout the process, adherent clusters can easily become detached by simply moving the flask; therefore, it is of utmost importance for the integrity of the culture system to absolutely avoid any movement of the flask or its content during the crucial aggregate-toMAN transition process. Once hNPC clusters have detached, they will immediately merge with any other suspension aggregates they come into contact with (via integrins and secreted ECM proteins), thereby perturbing the essential spacing component of adherent growth. Even removal of the flask to view under the microscope disrupts the culture setup and should be avoided. For the same reasons, it is not prudent to supplement growth factors during this time; therefore, MAN cultures are started in 40 ng/ml bFGF to account for rapid degradation and resultant mitogen loss over the first 48 hr.

4. After 3 to 4 days of untouched growth, gently move the flask from the incubator to the sterile hood. 5. Slowly tilt the flask up to 90◦ , then slowly rock backwards so that the flask is now upside-down, the CM is now facing downward on the top of the flask, and the cellular plane is facing upwards. You should be able to visibly identify exposed adherent clusters attached to the flask.

6. Aspirate all of the medium from the flask and, quickly but gently, add 8 ml fresh NB-B-27 complete medium containing 20 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin to the downward (noncellular) plane, being careful not to slosh medium onto the upper (cellular) plane, which would dislodge the lightly adherent cells. Do not allow the flask to dry out after the medium has been aspirated, as hNPC may begin to detach upon reintroduction of fresh medium to the culture.

7. In a reverse motion, rock the flask back slowly to its original position, paying careful attention as the medium re-covers the adherent cells. During this process, it is absolutely imperative to reintroduce the fresh medium in a slow fluid motion to minimize waves as the medium spreads across the flask. Any major fluctuations or tapping of the flask can easily dislodge the clusters from their equally spaced positions, threatening the overall integrity of the MAN. No matter how careful you may be, there will always be a small percentage of cells that either did not attach or have detached during the feeding process. These floating cells will either reattach or can be removed from the culture at the time of the next feeding.

8. After the medium has been changed, place the flask back into the incubator and repeat the process every 2 to 3 days as necessary to replenish growth factors (48-hr half-life) or replace metabolized medium (indicated by an orange acidic appearance). The literature and product datasheets support a general half-life for most of the growth factors used in this unit at 24 to 72 hr at 37o C in these medium formulations. The cells also utilize a large proportion, so we generally assume that the majority of the growth factors need to be replenished; therefore, we supplement according to the volume in the

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flask and adjust the concentration to the full concentration on the assumption that there is no growth factor remaining. As the cultures expand, it will become necessary to alter the percentage of CM exchanged. During the first week, 75% to 100% of the medium should be exchanged to account for metabolized nutrients while maintaining adequate paracrine conditioning. As cultures develop from week one onward, it will become essential to exchange 100% fresh NB-B-27 growth medium every 1 to 2 days to replenish the highly metabolized nutrient stores and remove toxic metabolic byproducts. CM does not need to be added back in this case, as the high density-to-volume ratio leads to quick paracrine conditioning, adequate for immediate sustained survival. Furthermore, these fully developed MAN cultures can be utilized for the collection of high-quality CM (Support Protocol 4). Over the next 2 to 3 weeks, MAN hNPC continue to proliferate and spread into a webbed culture, whereby adherent cellular islands will not only expand into each other but also proliferate in the vertical z dimension, creating the characteristic multilayer threedimensional appearance (Fig. 2D.3.1G-I). As the MAN matures, it will develop into a highly mitotic (75% to 85%) confluent culture. Although clusters will continue to merge, there will always be demarcated areas on the flask surface where no hNPC grow; therefore, these cultures never attain the classic two-dimensional monolayer morphology.

Feed Multilayer Adherent Network (MAN) MAN cultures offer many time and growth advantages over classic aggregate or suspension sphere assays. Care should be taken to minimize sloshing of medium or excessive vibration that will detach the fragile network of cells. The basic rule for ease of use with this system is to minimize mechanical stress, especially at the edges of the flask, which can easily loosen the outer edges of the MAN, exposing the undersurface and resulting in uplifting of the entire sheet of adherent progenitor cells. Although these adherent networks of cells appear to be stably anchored to the flask, it takes relatively little force to disrupt their fragile connections. Furthermore, once detached, the cells will remain adherent in their networks and organize into large clumps, floating or partially attached to the remaining sheet of cells, which may become necrotic if not dissociated in ample time. Any cellular debris and insoluble salt residues that may develop from prolonged culture are removed by the methods described below. 9. Slowly tilt the flask up to 90◦ and rock backwards so that the CM is facing downward on the top of the flask and the cellular plane is facing upwards. Carefully aspirate or collect conditioned medium See Support Protocol 4 for treatment of the conditioned medium.

10. Gently rinse the flask once with 8 ml DPBS (for 25-cm2 flask) or 12 ml DPBS (for 75-cm2 flask) by expelling DPBS onto the downward (noncellular) plane at low speed, being careful not to slosh liquid onto the upper (cellular) plane, which would dislodge lightly adherent hNPC. Do not allow the flask to dry out after DPBS has been aspirated, as hNPC will begin to detach upon reintroduction of fresh media to the culture.

11. In a reverse motion, rock the flask back slowly to its original culture position, paying careful attention as the DPBS re-covers the adherent cells. During this process, it is absolutely imperative to reintroduce the fresh DPBS in a slow fluid motion to minimize mechanical fluctuations as it spreads across the flask Long-Term MAN Growth and Characterization of NPCs

12. Repeat steps 9 to 11, transferring 8 to 10 ml (for 25-cm2 flask) or 15 to 20 ml (for 75-cm2 flask) fresh NB-B27 complete medium (containing 20 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin) to each flask. Slowly move the flask to a humidified incubator at 37◦ C, 5% CO2.

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Dissociate Multilayer Adherent Network (MAN) For extensive discussion of factors that are critical to the dissociation (passaging) of MAN hNPC cultures, see Critical Parameters and Troubleshooting. 13. When cultures are ready for passaging (see Critical Parameters and Troubleshooting), slowly tilt the flask upwards to 90◦ , then rock backwards so that the CM is facing downward on the top of the flask and the cellular plane is facing upwards. Aspirate or collect conditioned medium. See Support Protocol 4 for treatment of the conditioned medium.

14. Gently rinse the flask once with 8 ml CMF-DPBS (for 25-cm2 flask) or 15 ml CMFDPBS (for 75-cm2 flask) by expelling CMF-DPBS onto the downward (noncellular) plane, being careful not to slosh CMF-DPBS onto the upper (cellular) plane, which would dislodge lightly adherent cells. Do not allow the flask to dry out after medium has been aspirated, as hNPC will begin to detach upon reintroduction of fresh liquids to the culture.

15. In a reverse motion, rock the flask back slowly to its original position, paying careful attention as the CMF-DPBS re-covers the adherent cells. Repeat step 13 and aspirate. During this process, it is absolutely imperative to reintroduce the CMF-DPBS in a slow fluid motion to minimize mechanical fluctuations as it spreads across the flask.

16. Gently add 3 to 5 ml (for 25-cm2 flask) or 7 to 10 ml (for 75-cm2 flask) of Accutase (prewarmed to 37◦ C, 10 min before use) to flask without disrupting the integrity of the cellular sheet (as described for CMF-DPBS rinse in steps 13 to 15). 17. Carefully transfer the flask into a 37◦ C, 5% CO2 humidified incubator for 3 to 5 min (depending on density), minimizing any significant motion that will release the multilayer adherent network prematurely. The key to the successful dissociation of a MAN culture relies on learning to recognize the following properties throughout the incubation in dissociation agent. a. As the enzyme initially begins to break down cell-cell contacts, the adherent culture releases from the plastic dish from the outside in. Generally speaking, the outermost edges of the network will flap up and off of the dish, generating an organized sheet that eventually releases from the plastic dish below. If the dish is prematurely interrupted during this incubation process by moving the flask or sloshing the Accutase solution, the precise coordinated lifting of the multilayer adherent network is disturbed and subsequently leads to breakdown of the intact sheet of cells. Inadvertent disruption of the intact sheet can lead to gross clumping and compromise the integrity of cells as they dissociate. b. In addition, prolonged exposure to enzymes can puncture the cell membrane and render hNPC extremely vulnerable to mechanical shearing, resulting in lysis and release of DNA into the cell suspension. The results of enzyme overexposure are visibly apparent, as evidenced by increased viscosity of the cell suspension accompanied by discernibly large floating aggregates. These aggregates have a propensity to float to the top of the cell suspension and are characterized by their sticky, slimy properties that render them problematic in culture as they accrue and amass live cells on the surface. As the aggregates continue to bind live hNPC, they become heavier and eventually fall by gravity from the top of suspension to the bottom, thus allowing for removal from the remaining population. The overall result of enzyme overexposure is decreased hNPC recovery; therefore, it is imperative to time the enzymatic process and visually inspect the flask after 3 to 3.5 min, to monitor the dissociation progress closely. c. During the 3- to 5-min incubation process, the MAN layer will gradually detach completely from the underlying flask, effectively shrinking into an intact rectangular sheet, resembling a miniature compacted version of the original MAN. The exact timing for completion of this process is variable, but should be minimized to account for overexposure. In general, the entire sheet should be detached and shrunken into the center of the flask

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for at least 1 to 3 min before the desired stage of dissociation is attained. Note that this is an extremely time-sensitive process. Lesser incubation times will result in incomplete dissociation of larger hNPC clusters, requiring additional cycles, ultimately leading to increased clumping and subsequent cell death.

18. After 3.5 to 5 min, when the MAN displays the above characteristics, gently transfer the flask to a sterile hood, paying special care to retain the free-floating cellular sheet in its intact form for easy removal. The intact sheet is extremely fragile and will most likely begin to dissociate as the flask is moved. Try to retain the sheet in as many large pieces as possible. Furthermore, lowerdensity cultures will not retain the structural integrity that their mature MAN counterparts display.

19. Carefully tilt the flask so that the sheet of cells aggregates to the bottom corner of the flask with gravity. With a 5-ml pipet, carefully suck up the concentrated network of cells in 1 to 3 ml of the Accutase solution and transfer to a 15-ml conical tube. It should be possible to reclaim the cells into a small volume without extensive single cell dissociation or disruption of the cellular sheet. The remaining Accutase should appear clear and may contain a few smaller cell clusters.

20. Gently triturate contents of the conical tube with a 5-ml pipet attached to a pipetting aid (e.g., Drummond) on medium speed (five to seven times) to break the cell suspension into smaller floating cellular aggregates. Be very careful not to over-triturate, as the cell suspension is extremely fragile at this stage.

21. Using the same 5-ml pipet, immediately triturate the remaining contents of the flask to break up remaining clusters, gently but thoroughly, paying extra attention to the removal of adherent hNPC at the edges of the flask where they tend to attach preferentially and with increased strength. Transfer the contents of the flask to the previous conical tube. 22. Continue trituration of hNPC inside the conical tube to break the cells up into smaller clusters by gently expelling the cell suspension at a 45◦ angle against the wall of the conical tube at medium speed (8 to 10 times). 23. If necessary, recap the conical tube and incubate in a 37◦ C water bath for 1 to 2 min more with constant swirling to avoid clumping of aggregates at the bottom of the tube and reduce accumulation of sticky DNA from lysed cells. It is very important to ensure the hNPC do not aggregate and begin clumping during the dissociation process; therefore, care should always be taken to continuously swirl or triturate the cells during steps 20 to 23.

24. Using a 1000-μl extended-length pipet tip with a standard automatic pipettor set to 750 μl, slowly triturate hNPC suspension at a 45◦ angle against the wall of the conical tube at a consistent rate. Excessive or high-rate trituration against the plastic wall is not well tolerated at this stage. We recommend slow to medium trituration at a position near, but not touching directly against the wall of the conical tube (five to ten times or until large clumps are no longer visible and the dissociated solution has a homogenous milky and sandy appearance). Ideally passaged cultures will be fully dissociated into single cells, >95% viable, and free of floating aggregates if the time of initial Accutase exposure was within the correct window (step 17), cells are not allowed to aggregate, and trituration remains moderate and minimal. Long-Term MAN Growth and Characterization of NPCs

Cell clusters will readily stick to the meniscus (∼750-μl line) of the pipet tip.

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25. To recover cells that have stuck to the meniscus, reset the plunger from 750 μl to 1000 μl (with tip remaining intact). Rinse the 1000-μl tip once with 1000 μl NB-B-27 complete medium to dislodge residual clusters, and transfer the contents to a new 15-ml conical tube containing 10 ml fresh NB-B-27 complete medium containing 20 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin (prewarmed to 37◦ C) to inactivate the reaction. 26. Inactivate fully dissociated preparation from step 24 by adding it to the 10 ml medium in the conical tube from step 25. Variability in hNPC culture densities and morphology will dictate the specific timing and rate of dissociation for each culture. As a result, it is often the case that a small percentage of undissociated cell clusters remain and require a second round of enzymatic treatment, while the majority of cells are fully dissociated and ready to be inactivated and released from enzymatic shock.

27. To process partially dissociated cell suspensions, place the conical tube vertically for 1 to 2 min until the visible cellular clusters have settled by gravity to the bottom. Carefully transfer the top portion of supernatant containing dissociated cells to the previously inactivated cell suspension. To the remainder of undissociated hNPC, add 1 ml fresh prewarmed Accutase, triturate twice, and repeat steps 24 to 26. 28. Transfer the appropriately dissociated cell suspension to the previously inactivated 10 ml hNPC suspension from step 26. In rare cases, some clusters may remain after the second round of dissociation (often seen in necrosis) and are considered behaviorally abnormal and subsequently discarded. CAUTION: Overexposure to any dissociating agent will cause significant cell death and deter growth from lysed hNPC. The solution will become more viscous when this occurs. Thus, the procedure should be optimized to break up the cell clusters, while minimizing the amount of time in the dissociation agent. Generally, the larger the flask, the more dissociation agent that will be needed, which means more cell death and greater difficulty in controlling the timing of the process. We recommend 25-cm2 or 75-cm2 flasks for optimal conditions.

29. Centrifuge the cell suspension for 4 min at 400 × g, room temperature. Carefully aspirate the supernatant. Adherent cultures exhibit a highly branched, polarized cellular morphology, and unfortunately many of these delicate processes are cleaved by dissociating agents and mechanical stress, resulting in a greater amount of cellular debris. As a result, an additional rinse and centrifugation with 10 ml of either CMF-DPBS or Neurobasal medium (Invitrogen) is recommended to remove any problematic residual debris.

30. Resuspend the hNPC pellet in the conical tube with 1 ml fresh NB-B-27 complete medium using an extended-length 1000-μl pipettor and tip, gently triturating five to seven times to thoroughly liberate the cell pellet. 31. Count viable cells using a hemacytometer and trypan blue (UNIT 1C.3) for correct replating density. 32. After counting, add 8 ml CM (for a 1:2 dilution) to the conical tube, adjust for the desired final volume of fresh NB-B-27 complete medium to CM ratio accordingly (i.e., 8 ml fresh NB-B27 medium for 50% CM final), bring cells to desired density, and replate into new 25-cm2 flasks. In general, more concentrated splits survive and proliferate more effectively than their diluted counterparts. As a guideline, a 25-cm2 flask containing 1–3 × 106 cells is fed 25% to 50% CM, and 4 × 106 cells do not require CM as they quickly condition the medium due to high density.

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33. Add bFGF and LIF to achieve a final concentration of 20 ng/ml and 10 ng/ml, respectively. Gently swirl contents of flask horizontally to evenly disperse hNPC and place in a humidified incubator at 37◦ C, 5% CO2 . Subsequent culturing methods will depend on the density of cells plated and method for further expansion.

34. Passage MAN cultures. Typically, the growth parameters of hNPC MAN cultures dictate passaging once every 1 to 2 months depending on the original plating density and desired confluency. We typically split MAN cultures at a 1:2 dilution for 3–4 × 106 cells/25-cm2 flask of mature 65% to 75% confluent culture, or 1:4 for 5–10 × 106 cells/25-cm2 flask of very mature 80% to 90% confluent extremely high-density 2-month-old cultures, as they contain many more cells per flask than a typical aggregate culture where high density cannot be achieved at the cost of fusion, large globular aggregate formation, and ensuing necrosis. We consider the above modifications of the enzymatic process, specifically the precisely timed controlled release of the entire MAN as an intact sheet, to be one of the key components of successful passaging and subsequent expansion of hNPC using this assay. Consistent high viability and overall health of the resultant hNPC preparations coupled with the intrinsic quantitative qualities of the assay (i.e., increased population doubling rate, apparent increase in proliferation capacity for >100 passages without senescence or decease in rate of replication, and decreased cost in consumables and personal time) all mark the overall utility and advantages for employing the MAN assay to obtain long-term expansion of large quantities of undifferentiated hNPC. MAN cultures can also be processed by traditional methods used for aggregate cultures. Simply triturate adherent cells thoroughly from the flask and proceed as described for aggregate cultures (Alternate Protocol 1). It should be noted that enzymatic dissociation times will be greatly enhanced, requiring multiple rounds of gravity-based cluster separation, enzymatic treatment, and subsequent centrifugation cycles. Unfortunately, this procedure results in significant cell death (60% to 70% viability) in even the most skilled hands, and should only be employed when cells are accidentally detached by mechanical force. In these cases, a second rinse and centrifugation step should be added prior to final plating.

SUPPORT PROTOCOL 1

DERIVATION OF HUMAN FETAL NEURAL STEM/PRECURSOR CELLS Fetal spatial features and their specific neuroanatomical coordinates are used to determine the cadaver’s specific stage of CNS development and dictate the exact location for tissue dissection. Proficiency in fetal neuroanatomy is essential for efficient assessment and subsequent resection of specified CNS regions. We, along with others, have described various methods for the derivation of hNPC. Here, we detail the methodology we have successfully employed to isolate and expand fetal forebrain periventricular zone human NPC. NOTE: Use of human fetal cadaveric CNS must follow all safety and bioethical guidelines, including but not limited to full informed consent, IRB approval, and strict adherence to all state and federally mandated laws and guidelines for the ethical use and treatment of patients or specimens derived thereof (also see APPENDIX 1A).

Long-Term MAN Growth and Characterization of NPCs

NOTE: Perform all procedures aseptically in a sterile Biosafety Level 2 hood. Sterilize all surgical tools in a hot bead sterilizer or autoclave (121◦ C, 2 hr), or by gas sterilization. During the procedure, place all of the tools in fresh 70% ethanol when not in use. Immediately following removal from ethanol, briefly rinse twice in fresh sterile DPBS (Mediatech, cat. no. 21-031-CM).

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Materials Fetal tissue 10% (v/v) formalin (optional) Enzymes for tissue dissociation (optional): e.g., Accutase, trypsin-EDTA, PPD (papain-protease-DNase I) Fetal bovine serum (FBS; optional) NB-B-27 complete medium (see recipe) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) Leukemia inhibitory factor (LIF; Millipore, cat. no. LIF1010) Normocin (InvivoGEN, cat. no. ant-nr-1) Epidermal growth factor (EGF; Millipore, cat. no. 01-107) Surgical equipment, including scalpel, sterile 15-ml conical tubes Battery-powered pipetting aid (e.g., Drummond Pipet-Aid XP) Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) Isolate and digest human fetal periventricular zone 1. Stage the fetus using neuroanatomical coordinates, open the head cavity, and remove the brain. 2. Cut sagittally across the midline to separate the cerebral hemispheres then cut again coronally from frontal to occipital poles. 3. Select the brain slice containing the region of interest for dissociation. Optional: Fix the remaining tissue in 10% (v/v) formalin for a more extensive neuropathological examination.

4. Carefully scrape the ventricular wall and adjacent subventricular zone region from the forebrain section with a surgical scalpel. Delicately mince the dissected tissue into small pieces with the scalpel blade. 5. Transfer the tissue pieces into a 15-ml sterile conical tube that contains 6 ml cold NB-B-27 medium, 20 ng/ml bFGF, 20 ng/ml EGF, and 4 μl/ml Normocin. 6. Place the conical tube vertically and allow the tissue to pellet by gravity (1 to 2 min), aspirate supernatant carefully, and rinse three times, each time with 8 ml cold medium. 7. After final rinse, resuspend the tissue in 8 ml cold medium. 8. Gently triturate the fetal tissue suspension (10 to 15 times) with a 5-ml pipet attached to a pipetting aid (e.g., Drummond Pipet-Aid XP) at medium speed against the wall of the 15-ml conical tube to further dissociate the tissue into a homogenous milky solution. The cell suspension will contain both single cells and a few small cellular clumps. Try to avoid introducing air bubbles during the trituration process. It is important that the primary tissue not be overzealously digested into a single-cell suspension, due to the subsequent damage incurred by mechanical stress on the progenitor fraction. CNS tissue from young fetal brains is softer than that from fully developed myelinated adult brains; therefore, later-stage CNS preparations include the addition of an enzymatic agent such as Accutase, trypsin-EDTA, papain-protease-DNase I (PPD), dispase, or any commercially available reagent, according to the manufacturer’s instructions, to efficiently dissociate primary cultures before their initial plating. In general, enzymatic fetal tissue dissociation averages ∼5 to 10 min, while adult tissue can take upwards of 45 to 90 min to generate the desired breakdown of brain tissue.

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9. Remove large undissociated tissue bits remaining after the initial trituration by allowing them to settle by gravity (2 to 3 min), then collect the suspension of cells in the upper supernatant. Dissociate remaining undissociated cell clumps again as in the steps above and pool together with the originally dissociated cell suspension. 10. Inactivate enzymatic preparations by diluting them 1:5 in fresh prewarmed NB-B27 medium and centrifuge for 5 min at 400 × g, room temperature. Remove supernatant and retain pellet.

Establish primary hNPC cultures 11. Following primary dissociation, bring the cell suspension to working volume in 8 ml pre-warmed NB-B-27 medium with 20 ng/ml bFGF, 20 ng/ml EGF, and 4 μl/ml Normocin at a final density of 1 × 105 cells/cm2 in one 25-cm2 flask and place in a humidified 5% CO2 incubator at 37◦ C. Primary cultures plated onto tissue culture treated flasks will generally produce mixed aggregate and adherent cultures. Primary cell suspensions may also be plated onto fibronectin-coated tissue culture–treated flasks for monolayer-like (two-dimensional) adherent cultures.

12. Determine cell viability using either the propidium iodide or trypan blue exclusion assay and a hemacytometer (UNIT 1C.3). Sticky cellular debris and small undissociated neural clumps may make this process difficult initially.

13. Optional: Add 0.1% to 1% (v/v) fetal bovine serum (FBS) at the time of initial derivation to enhance initial NPC expansion efficiency, promote adhesion, and decrease overall cell death with a relatively low risk of differentiation. CAUTION: Using FBS may introduce unwanted variability. Serum components are removed after a short period of time and replaced with a defined, serum-free medium so as not to potentiate long-term side effects on primary hNPC cultures. In some cases, it is desired that newly derived stem cell lines be established utilizing serum-free protocols so as not to introduce animal proteins into culture.

14. Incubate cells. At a time point 12 to 48 hr after plating, rinse any serum-containing cultures twice with 10 ml DPBS and transfer cultures to serum-free conditions in NB-B-27 medium containing 20 ng/ml EGF, 20 ng/ml bFGF, 10 ng/ml LIF, and 4 μl/ml Normocin. Continue incubation. 15. At a time point 3 to 4 days after the primary plating, supplement cultures by carefully removing the top half of medium from each flask, termed conditioned medium (CM), and replace with fresh NB-B-27 complete medium containing 40 ng/ml EGF, 40 ng/ml bFGF, 20 ng/ml LIF, and 8 μl/ml Normocin for the final working concentration of 20 ng/ml EGF, 20 ng/ml bFGF, 10 ng/ml LIF, and 4 μl/ml Normocin. These final concentrations are based on the assumption that the growth factors have been completely deleted by this point.

16. For more efficient recovery, remove the CM containing free-floating aggregates and small clumps of primary tissue and transfer the contents to a new flask. Triturate the cell suspension thoroughly to redissociate the remaining clumps, and supplement with fresh growth factors and antibiotics by the above procedure.

Long-Term MAN Growth and Characterization of NPCs

Alternatively, centrifuge suspension aggregates and debris for 3.5 min at 400 × g, aspirate, and either add the cells back to the original parent culture flask for further expansion or replate the primary cultures into 8 ml fresh pre-warmed NB-B27 complete medium containing 20 ng/ml EGF, 20 ng/ml bFGF, 10 ng/ml LIF, and 4 μl/ml Normocin.

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17. Repeat steps 15 and 16 throughout the first few weeks of primary culture. Initially, hNPC will proliferate throughout the flask as a mixture of adherent and freefloating aggregates and can be detached from the culturing vessel through repeated trituration. We stress the inclusion of adherent monolayer-like hNPC within primary cultures during the initial hNPC expansion stage. As cultures mature, adherent hNPC cultures may also spontaneously give rise to a few spherical balls. These aggregates detach from the initial colony and continue to expand and self-renew as free-floating suspension cultures as well.

18. After several weeks, select the hNPC cultures that proliferate in a morphologically relevant manner and dissociate into single-cell suspensions or small clumps (3 to 8 cells/clump) with Accutase or cell dissociation buffer (CDB)/cellstripper. Dissociate when cellular aggregates are larger than 12 to 15 cells in diameter and can no longer be mechanically separated by simple trituration or when adherent cultures become greater than 75% confluent. Pool both adherent and free-floating cells and discard any remaining large clumps that do not readily dissociate. 19. Replate hNPC at a 1:1 or 1:2 ratio as either multilayer adherent aggregates or as suspension aggregates in NB-B-27 complete medium, 20 ng/ml EGF, 20 ng/ml bFGF, 10 ng/mL LIF, and 4 μg/ml Normocin for 2 more weeks. 20. Exchange one-half of the culture medium as described in step 15 every 2 to 3 days to replenish growth factors and antibiotics. Dissociate and replate cultures (1:1 or 1:2) once per week or as necessary. After 2 weeks, exclude LIF and EGF for mitogen selection.

Mitogen-select primary hNPC cultures After 2 to 4 weeks of primary expansion, undifferentiated hNPC colonies will proliferate and establish a healthy culture of precursors. At this point, successful cultures are subjected to a 10-week sequential growth factor selection process utilizing parameters of growth rather than markers alone to select for the proliferative EGF/FGF responsive population of cells. 21. Expand hNPC as a mixed population of both adherent clusters and free-floating aggregates in NB-B-27 complete medium containing 20 ng/ml bFGF alone (and 2 μl/ml Normocin) for 2 weeks with (1:1 or 1:2) dissociation once per week throughout the selection process as dictated by size exclusion and morphological parameters described above in step 18. 22. After 2 weeks, omit bFGF and supplement the medium with 20 ng/ml EGF alone (and 2 μl/ml Normocin) for 2 weeks. 23. Maintain the bFGF/EGF 2-week rotation schedule for two to three sequential rounds (10 weeks) and complete after the final bFGF-alone cycle. 24. After the final selection process, a few primary hNSC/hNPC cultures will continue to proliferate and display appropriate morphology; dissociate these cultures and pool together into NB-B27 complete medium containing 10 ng/ml LIF, for a final hNPC complete basal maintenance medium composed of NB-B-27 growth medium containing 20 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin for secondary hNPC expansion.

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ALTERNATE PROTOCOL 1

FEEDING AND DISSOCIATION OF LIGHTLY ADHERENT AGGREGATE CULTURES Human NPC can also be successfully expanded by traditional aggregate culture without extracellular matrices when plated at a density no less than 1–2 × 106 cells/25 cm2 . At a time point 2 to 3 days after dissociation, small (4- to 8-cell) clusters form and will proliferate as both suspension aggregates and lightly adherent clusters. Cultures are fed fresh medium and growth factors two to three times per week, depending on the specific density and metabolic capacity. Approximately every 2 days, cellular aggregates will project lightly adherent processes onto the plastic surface. These clusters are triturated gently with a 5-ml pipettor and supplemented with growth factors for a final concentration of 20 ng/ml bFGF and 10 ng/ml LIF. Detailed procedures can be found elsewhere (Wakeman et al., 2009). Lightly adherent cellular clusters are enzymatically passaged with Accutase when they grow larger than 12 to 15 cells (100- to 150-μm) in diameter or can no longer be readily broken apart mechanically by gentle trituration (approximately once per week).

Materials Human NPC growing in 25-cm2 flasks (Support Protocol 1) NB-B27 complete medium (see recipe) Accutase (Millipore, cat. no. SCR005) or Cell Dissociation Buffer (Invitrogen, cat. no. 13150-016) Battery-powered pipetting aid (e.g., Drummond Pipet-Aid XP) 15-ml conical tubes Centrifuge Pipettors with extended-length pipet tips 1. Triturate the contents (minimizing bubbles) of a 25-cm2 flask of human NPC gently eight to ten times with a 5-ml pipet attached to a pipetting aid (slow speed) to detach lightly adherent cellular clusters from the plastic surface. Transfer the contents of the flask to a 15-ml conical tube. Rinse the flask with 2 ml fresh prewarmed NB-B-27 growth media to collect any residual hNPC and transfer to previous conical tube. Triturate the entire surface by tilting accordingly, paying careful attention the corners of the flask, where cells tend to preferentially adhere.

2. Centrifuge 3 to 4 min at 400 × g, room temperature. Remove supernatant from conical tube and filter the conditioned medium (CM). Treatment of the conditioned medium is described in Support Protocol 4.

3. With a 1000-μl pipettor and extended-length pipet tip, dropwise add 750 μl Accutase to the conical tube and carefully triturate the hNPC pellet three to five times lightly against the wall of the tube to dislodge the cells. The pipettor should never touch the side of the conical tube while pulling the solution up and down into the tip. Extended-length pipet tips allow for easier access into the conical tube and reduce the chance of contamination.

4. Place the conical tube into a 37◦ C water bath and incubate 3 to 5 min with constant swirling to avoid settling and clumping of hNPC. 5. Proceed to steps 24 to 33 in Basic Protocol 1. Long-Term MAN Growth and Characterization of NPCs

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GROWING hNPC IN MAN MEMBRANE SYSTEM (MMS) To better accommodate analysis and manipulation of hNPC, we have adapted the MAN culture system for growth on transferable semi-porous membrane inserts, termed the MAN membrane system (MMS). MMS cultures offer an extensive variety of choices in experimental design. Perhaps the most beneficial feature of the MMS is the ease of manipulation that the system offers, as basket inserts are movable and amenable to a plethora of biochemical, growth, and cytokine migration assays. Cells are always easily accessible and can be dissociated or removed from the membrane using the same procedure as the MAN assay. For this reason, we prefer utilizing MMS cultures during lentiviral infections (example can be found in Alternate Protocol 4). The MMS baskets can easily be rinsed and moved from clean well to clean well by simply removing the insert. As a result, the proliferative network of hNPC never has to be disrupted, increasing the infection efficiency as well as the viability of cells post-infection.

ALTERNATE PROTOCOL 2

Materials NB-B-27 complete medium (see recipe) Leukemia inhibitor factor (LIF; Millipore, cat. no. LIF1010) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) Normocin (InvivoGEN, cat. no. ant-nr-1) Freshly dissociated hNPC or small aggregates (Support Protocol 1) 6-well tissue culture plates Forceps, sterile 1.0 to 0.1-μm hanging basket transmembrane cell culture insert (Corning) 1. Add 3 ml NB-B27 complete medium containing 40 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin, prewarmed to 37◦ C, to each well of a tissue culture treated 6-well culture plate. The concentration of bFGF is increased to due to the additional incubation time necessary to induce MAN growth characteristics.

2. Using a sterile forceps, insert one 1.0 to 0.1 μm hanging basket transmembrane cell culture insert into each well. We utilize polyethylene terephthalate membranes because they offer great optical properties as well as excellent adherence. In addition, we find that hNPC can spontaneously migrate through any pore larger than 1.0 μm, albeit in low proportions.

3. Transfer 1.0 × 105 freshly dissociated hNPC or small aggregates in 2.5 ml NB-B-27 complete medium per basket insert. 4. To allow for adequate attachment, culture undisturbed at 37◦ C in a humidified 5% CO2 incubator for 72 to 96 hr to induce MAN features. The porous membrane allows hNPC to efficiently attach and often confers adherence more quickly than standard tissue culture plastic. In addition, once the MAN has established adherence, medium can be safely aspirated from the lower chamber without disrupting the fragile network of hNPC in the upper basket insert.

5. Every 2 days for 2 to 3 weeks of culture, replace 100% of the medium in the lower basket as well as 50% of the CM in the upper portion of the basket insert with fresh NB-B-27 complete medium containing 20 ng/ml bFGF, 10 ng/ml LIF, and 2 μl/ml Normocin. Remove medium from the lower chamber first, followed by the basket; otherwise the basket will bob up and down and detach the fragile adherent network of cells. Slowly aspirate from the upper meniscus of medium so as not to disrupt the MAN when replacing medium from the basket. Medium will begin to slowly drip by gravity through the basket to the lower chamber. Although medium freely moves by gravity from upper to lower chamber

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when the lower chamber is empty, the same mixing effect does not occur when both chambers are full. As cultures mature, you will notice that the medium within the upper basket becomes metabolized and does not necessarily fully equilibrate with the lower chamber (i.e., upper-basket-insert medium will be orange and lower-chamber medium will be red). This suggests that medium does not efficiently mix between the chambers; therefore, it is imperative to change the upper basket medium as well to remove toxic metabolites and any dead cellular debris that may accumulate with normal growth. During this period, the MAN will become established and develop into a robust multilayer webbed interfaced network, creating classic MAN three-dimensional honeycomb structures composed of healthy, highly proliferative, multipotent, migratory hNSCs at a density of ∼2 × 106 cells/insert by 2 to 3 weeks. The overall rate of cell proliferation in our MMS system appears to drastically increase the replicative capacity we have seen previously in the HFB-2050 fetal hNPC line utilizing the classic neurosphere assay. In addition, we have seen no change in proliferative capacity over extensive periods of time or at high passage number (>60) when utilizing these methods. We have noted that the cellular dynamics of this system are highly dependent on the presence of LIF in the culture medium owing to an unknown mechanism most likely not related to protection of telomeres. Removal of LIF results in a situation highly mimicking that of MAN cultures on traditional tissue culture plastic; moreover, MMS cultures are phenotypically indistinguishable from MAN cultures, suggesting that the porous membrane does not confer any additional adhesion properties. We believe cells adhere and may proliferate at an elevated rate due to the additional trophic support and nutrient exchange conferred through the semi-porous membrane underneath the network of hNPC. Bidirectional nutrient exchange allows hNPC cultures to thrive from both sides, creating an ideal environment for three-dimensional proliferation within a two-dimensional lattice. SUPPORT PROTOCOL 2

CRYOPRESERVATION OF hNPC Cryopreservation of early-passage batched hNPC populations allows the researcher to thaw and expand aliquots of cells at a later point in time for experimental replication or to allow outside investigators to compare and contrast them with their own independently derived precursor lines. We freeze aliquots of hNPC in large batches every five population doublings to ensure that low-passage cells will be available in adequate numbers for extended studies. We always freeze hNPC as small multicellular aggregates versus single cells to increase recovery post-thaw. For the best results, hNPC are dissociated into single cells 48 to 72 hr before freezing, producing small (8 to 16 cells/cluster) to medium (16 to 32 cells/cluster) size clusters. During this short period of growth, 10% to 20% of hNPC may actively divide; however, this proliferation is offset by the 10% to 20% cellular death attributed to freeze/thaw cycling. Therefore, the number of hNSC originally dissociated is roughly equivalent to the number of cells that survive the entire freeze/thaw process. Freezing medium (see Reagents and Solutions) is made fresh at 4◦ C on wet ice at the time of use.

Materials 70% ethanol Cultures of hNPC grown in 25-cm2 flasks dissociated 48 to 72 hr earlier (Support Protocol 1) NB-B-27 complete medium (see recipe) hNPC freezing medium (see recipe) Liquid N2

Long-Term MAN Growth and Characterization of NPCs

1.8-ml cryovials (Nunc, cat. no. 377267) and labels 15- and 50-ml conical tubes Battery-powered pipetting aid (e.g., Drummond Pipet-Aid XP) Controlled-rate freezing device (e.g., “Mr. Frosty”; Nalgene) Liquid N2 tank

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1. Print labels for each cryovial, recording the date, passage number, and any other information pertinent for proper identification at later time of use. Inside the sterile hood, place a label around each cryovial and wipe thoroughly with 70% ethanol. Use an ink that will not fade in liquid nitrogen or upon exposure to alcohol.

2. Close the hood and turn UV light on for 2 hr. Open the hood and allow it to equilibrate for 15 min. Loosen the caps from the cryovials to allow for easy access. 3. Transfer a 25-cm2 flask of hNPCs to the hood. Gently dislodge any adherent cellular clusters from the flask by mechanical trituration [using a 5-ml pipet attached to a pipetting aid (e.g., Drummond) at high speed] of medium. Transfer the contents of the flask into a 15-ml conical centrifuge tube(s). Only freeze cultures that were dissociated 48 to 72 hr earlier.

4. Rinse the flask with 4 ml fresh pre-warmed NB-B-27 complete medium containing 2 μl/ml Normocin and add to the previous 15-ml conical tube. Centrifuge for 3.5 to 4 min at 400 × g, room temperature, to pellet hNPC. Aspirate the supernatant. Alternatively, transfer conditioned media (CM) supernatant to a conical tube and process (Support Protocol 4)

5. Gently resuspend the cell pellet by trituration with cold freezing medium (1 ml/1.8 ml cryovial). We generally freeze at a concentration of 1–3 × 106 cells/ml for medium to small clusters, respectively. Once the hNPC have been resuspended into freezing medium, the preparation process should be completed as quickly as possible to reduce the amount of time hNPC are exposed to the osmotic shock of DMSO. Depending on the density of the culture, one generally freezes four vials per 25-cm2 flask.

6. Evenly distribute the hNPC suspension among the sterile cryovials (at 1 ml/vial) and transfer the vials to a controlled-rate freezing device to cool the hNPC at ∼1◦ C/min. Human NPC clusters will quickly fall by gravity to the bottom of the cryovial; therefore it is best to freeze at maximum 10 to 15 vials at one time. Minimizing time and subsequent clumping of cells at the bottom of the each vial will dramatically increase the thaw efficiency. The ideal freezing duration occurs at a slow rate to reduce shock from crystallization and subsequent shearing.

7. Immediately place the freezing chamber in a −80◦ C freezer for 18 to 24 hr, then transfer cryovials to a liquid nitrogen tank or to a −140◦ C freezer for long-term storage. We have successfully thawed viable cells after over 10 years in storage using these methods.

THAWING CRYOPRESERVED hNPC During the freeze-thaw process, many hNPC will either die or differentiate, yielding ∼10% to 30% or ∼70% to 90% hNPC survival for single cells (Fig. 2D.3.3A) or small cellular clusters (Fig. 2D.3.3B), respectively. Freshly thawed hNPC are extremely fragile and highly susceptible to mechanical shear forces; therefore, careful processing of hNPC is essential for high-viability thaws and sustained expansion. In addition, it can take several weeks (post-thaw) to expand and amass a usable number of proliferative hNPC for subsequent experimentation. We generally utilize conditioned medium (CM) from thriving cultures to increase the rate of initial expansion, as it contains potent paracrine signaling molecules that stabilize and jump-start freshly thawed cultures. Always thaw small hNPC clusters (dissociated 48 to 72 hr before freezing) at a 2:1 or 1:1 ratio into the same volume/surface area as (or lesser than) the pre-freeze culture. Careful dilution of DMSO, gentle handling, and minimization of the duration of the thawing time are critical. Current Protocols in Stem Cell Biology

SUPPORT PROTOCOL 3

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Figure 2D.3.3 Cryopreservation and thawing of hNSC. Human NSC (HFB-2050) were dissociated into single cells just prior to cryopreservation (A), resulting in lower post-thaw yield. (B) When thawed as small aggregate clusters (arrow), imaged here at 12 hr post-thaw, fewer cells do not survive the freeze-thaw cycle (arrowhead).

Materials Frozen hNPC in 1.8-ml cryovials (Support Protocol 3) 70% ethanol Thaw medium: 50% (v/v) conditioned medium (Support Protocol 4)/50% (v/v) NB-B-27 complete medium (see recipe) Thaw medium (see above) containing 10 ng/ml leukemia inhibitory factor (LIF; Millipore, cat. no. LIF1010) and 20 ng/ml basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) 15-ml conical tubes 25-cm2 tissue culture flasks (non-ECM-coated) 1. Remove frozen hNPC vials from liquid nitrogen and place onto dry ice. CAUTION: Wear appropriate face and hand protection to protect from explosion of frozen vials.

2. Thaw one to two vials of frozen hNPC (1 to 2 ml) quickly with constant shaking until ice is almost cleared (∼60 to 90 sec) in a 37◦ C water bath. Rinse exterior of cryovial thoroughly with 70% ethanol and place into sterile tissue culture hood. 3. Carefully open the vial to release any built-up pressure, gently triturate the cell suspension twice by pipetting up and down with a 1000-μl pipettor/pipet tip to resuspend the cells, and immediately transfer the hNPC suspension into a 15-ml centrifuge tube containing 1 ml cold thaw medium. Excessive trituration at this point will induce significant cell death.

4. Dropwise, add approximately 6 ml cold thaw medium to dilute DMSO from freezing medium. Long-Term MAN Growth and Characterization of NPCs

As with cryopreservation, it is essential to transfer cells gently but quickly into cold medium, because extended incubation in DMSO will destroy hNSC by osmotic shock.

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5. Rinse the cryovial once with 1 ml cold thaw medium and transfer to the 15-ml conical tube. 6. Centrifuge the 15-ml conical tube 3.5 to 4 min at 400 × g, room temperature. Aspirate supernatant. Alternatively, the tube is placed vertically in an incubator at 37◦ C for 30 min to 2 hr. This step allows the cell clusters to settle by gravity but is not conducive to differentiation. Larger cell clusters will require less time to equilibrate to the bottom of the tube. The mixture of freezing and feeding medium is then centrifuged at 400 × g for only 1 to 2 min and the supernatant is safely aspirated. Aspiration of the medium without centrifugation will result in the loss of many cells.

7. Resuspend the hNPC pellet by gentle trituration with a 5-ml pipettor in 8 ml/25-cm2 flask of a mixture of 50% fresh NB-B27 complete medium and 50% CM plus 10 ng/ml LIF and 20 ng/ml bFGF, then replate onto 25-cm2 non-ECM-coated TC treated flasks. After 1 to 2 weeks in culture, the percentage of CM may be reduced from 50% to 25% and eventually 0% CM when nicely expanded adherent cultures are established.

PRESERVATION OF CONDITIONED MEDIUM Conditioned medium (CM) contains autocrine and paracrine effector molecules and can be utilized to enhance survival of hNPC during various procedural manipulations. For example, addition of CM to low-density cultures, to freshly thawed NPC, or as an aid in single-cell cloning can often be the key to a successful experiment. In an effort to collect relatively homogenous CM across samples, we apply a strict set of limitations on the quality of cultures that can be utilized to produce this paracrine-enriched basal medium supplement. Specifically, we only collect medium conditioned by healthy, highly proliferative, 65% to 75% confluent MAN cultures that have been grown in the medium for 20 to 24 hr. This procedure allows the cells to adequately secrete paracrine molecules into the medium without the cost of toxicity from metabolic breakdown of medium components over time.

SUPPORT PROTOCOL 4

Materials Human NPC MAN culture, 65% to 75% confluent (Basic Protocol 1) in 25-cm2 flask NB-B-27 complete medium (see recipe) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) Leukemia inhibitory factor (LIF; Millipore, cat. no. LIF1010) Normocin (InvivoGEN, cat. no. ant-nr-1) Acrodisc sterile syringe filter (0.2-μm; Pall, cat. no. 4433) 15-ml conical tubes 1. At a time point ∼20 to 24 hr prior to collection of CM, replace 100% of the medium in a 65% to 75% confluent MAN hNPC culture with 10 ml fresh NB-B-27 complete medium containing 10 ng/ml LIF, 20 ng/ml bFGF, and 2 μl/ml Normocin. 2. To harvest CM, carefully remove 10 ml CM from the 25-cm2 flask without dislodging and uplifting the fragile adherent network. Add 10 ml fresh media plus growth factors and resume incubation (it is possible to collect CM again from these cells as necessary). Slowly tilt the flask upside down for easier access to the medium.

3. Immediately filter 10 ml CM through a sterile 0.2-μm filter into a 15-ml conical tube.

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4. Add 20 ng/ml bFGF and 10 ng/ml LIF, mix by inversion, and freeze immediately at −20◦ C to limit degradation of paracrine signaling molecules. Prolonged exposure to environmental gas exchange, light, or room temperature conditions can rapidly degrade the CM, rendering it toxic or unbalanced as a salt solution.

5. To thaw CM, place the frozen tube at 4◦ C, overnight, to slowly melt the contents. If thawed tube contains any insoluble particles, discard immediately and thaw a new sample from a different batch.

6. Once the CM has thawed, prewarm to 37◦ C, mix according to the desired composition with fresh NB-B-27 complete medium, and supplement growth factors to the final appropriate concentrations. ALTERNATE PROTOCOL 3

REPLATING DISSOCIATED hNSC ON EXTRACELLULAR MATRIX (ECM) AS ADHERENT TWO-DIMENSIONAL MONOLAYER CULTURES In addition to the MAN assay described in detail here, hNPC can also be replated onto a variety of extracellular matrix (ECM) components at 1–2 × 106 cells/25-cm2 flask (maximum of 2–3 × 106 cells/25-cm2 flask) to induce attachment for more traditional two-dimensional, adherent monolayer growth parameters (Fig. 2D.3.4). As with MAN cultures, ECM attachment should not be utilized for low-density cultures where very few cell-cell contacts are present. The resulting cultures will likely become post-mitotic and differentiate prematurely. We prefer to expand primary hNPC lines without additional biological components, but we also recognize the utility and beneficial growth parameters that many ECM components confer in hNPC culture, especially when assaying and analyzing cells for migration and immunocytochemistry. That being said, not all ECM components are created equal, and each hNPC line will have its own particular characteristic adhesion properties. In our hands, hNPC tend to adhere to a variety of ECM proteins displaying a continuum for strength of adhesion—in order from weakest to strongest adhesion, fibronectin (human or mouse), laminin (human or mouse), Matrigel, collagen, and vitronectin. We recommend trying Millipore’s ECM cell culture optimization assay to determine the optimal ECM protein and concentration desired for the specific growth parameters chosen. In addition, a number of commercially available cell-binding enhancement solutions (Cell Bind) or specially scaffolded substrates (Cell Web, Corning) are also available, with a variety of binding properties to circumvent the use of biological attachment substrates. Furthermore, pre-coating flasks with electrostatically charged molecules such as poly-D-lysine or poly-L-ornithine in combination with extracellular matrix proteins provide a secondary level of support, often conferring an additional degree of adhesion. One warning is that poly-D-lysine should not be used for experiments involving electrophysiology, as it may interfere with ion-channel function.

Long-Term MAN Growth and Characterization of NPCs

In our hands, prolonged enhanced adhesion and exposure to matrix signaling molecules can have significant effects on hNPC phenotypic variation and related changes in cellular differentiation profile. For example, fibronectin supports a similar lightly adherent mode of growth to freshly dissociated MAN cultures on non-coated tissue culture–treated flasks, with the added benefit of slightly enhanced adhesion, quicker attachment, and higher rates of attachment. Laminin, likewise, retains many of the essential properties of the undifferentiated MAN, with the caveat that the initial adhesion is stronger, resulting in more flattened, monolayer-like, two-dimensional, multi-polar progenitor colonies. In slight contrast, Matrigel, a soluble basement membrane extract of the Engelbreth-HolmSwarm tumor, which is composed mainly of laminin as well as collagen IV, heparin sulfate proteoglycans, and entactin, but contains trace amounts of the platelet-derived growth factor (PDGF), nerve growth factor (NGF), insulin-like growth factor-1 (IGF-1),

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Figure 2D.3.4 Extracellular matrix confers two-dimensional monolayer phenotype in hNSC cultures. Human NSC (HFB2050) were plated onto tissue culture–treated flasks previously coated with a combination of poly-D-lysine and the extracellular matrix protein fibronectin. After 7 days in vitro, hNSC attain a similar composition and phenotype as MAN cultures, although they flatten and proliferate in a more two-dimensional manner in contrast to the three-dimensional architecture of MAN cultures (A-C). After 2 weeks, individual aggregate clusters are indistinguishable from each other, and begin to merge into a confluent layer of hNSC (D,E). In contrast to their MAN counterparts, these cultures will form a classic monolayer and lose their honeycomb appearance (F).

and TGF-β, supports exuberant growth of highly mitotic, extremely adherent, bipolar and multipolar neural precursors that will self-assemble into a highly dynamic neural niche (Watt and Hogan, 2000; Palmer, 2002; Wurmser et al., 2004; Lathia et al., 2007) composed of a heterogeneous population resembling type A, B, and C cells of the subventricular (SVZ) niche (D.R. Wakeman, unpub. observ.). Furthermore, substrates such as collagen IV and vitronectin bind hNPC, conferring an exceptional propensity for attachment, but typically at the cost of mass cellular differentiation. These findings introduce a secondary criticism of ECM components, in that ECM molecules naturally guide neuronal migration (Thomas et al., 1996; Murase and Horwitz, 2002, 2004; Labat-Robert and Robert, 2005; Flanagan et al., 2006; Hall et al., 2008) and are thought to play a critical role in differentiation of hNPC in vivo. As a result, culturing hNPC in the presence of these molecules in vitro may actually trigger primary differentiation of hNPC and an irreversible exit from the cell cycle. It is important, therefore, to choose an ECM accordingly and with respect to the specific assay of interest, as long-term cultures will adapt to their environment and may not continue to behave as true undifferentiated hNPC. We are comfortable with prolonged undifferentiated culture and expansion on either human fibronectin or human laminin (Ray et al., 1993; VicarioAbejon et al., 1995; Walsh et al., 2005; Flanagan et al., 2006; Ray and Gage, 2006; Hall et al., 2008) and temporary undifferentiated growth on Matrigel for 1 to 2 weeks.

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More adherent substrates such as vitronectin and collagen type IV are best utilized for differentiation assays. Specific brands and lots of ECM vary; therefore, proper testing is essential to determine individual growth parameters. It is worth noting that enzymatic lifting and dissociation of hNPC grown on strongly adherent ECM components generally require longer incubation times and often generate 10% to 20% cell death accordingly, due to the increased prevalence of fragile projections. For preparation of ECM substrates, see Support Protocol 5. SUPPORT PROTOCOL 5

PREPARATION OF EXTRACELLULAR MATRIX (ECM) SUBSTRATES Extracellular matrix can be applied to a variety of culture vessels. We recommend tissue culture flasks for expansion, 24-well plates with round glass coverslip well bottoms for differentiation, and multiwell chamber slides for routine immunocytochemical procedures. For enhanced ECM attachment, it is often useful or necessary to pre-charge the growth surface with poly-D-lysine or poly-L-ornithine.

Materials 100% ethanol Poly-D-lysine hydrobromide (Sigma, cat. no. P6407) or poly-L-ornithine (Sigma, cat. no. P4957) Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) 0.1% (w/v) fibronectin from human plasma (Sigma, cat. no. F0895) Laminin, human (0.5 mg/ml; Sigma, cat. no. L6274) or murine (Sigma, cat. no. L2020) Matrigel, growth factor–reduced (BD Bioscience, cat. no. 354230) Neurobasal medium (Invitrogen, cat. no. 21103049), cold Glass coverslips (Fisher, cat. no. NC970884) 24-well tissue culture plates Forceps, sterile 15-ml conical tubes Ziploc bag Prepare coverslipped plates 1. Wash coverslips thoroughly with 100% ethanol and autoclave prior to use. 2. Place one coverslip in each well of a 24-well plate with a sterile forceps.

Prepare poly-D-lysine/poly-L-ornithine solution 3. Create a stock solution containing 50 mg/ml of poly-D-lysine or poly-L-ornithine in water. Filter sterilize through a 0.22-μm Teflon filter, divide into aliquots, and store at –20◦ C. Charge substrate with poly-D-lysine/poly-L-ornithine 4. Coat the glass coverslips in the wells with sterile poly-D-lysine or poly-(L)-ornithine at 50 μg/ml. Incubate the solution overnight at 37◦ C. Aspirate. 5. Rinse five times with DPBS, 10 min each, to remove any toxic residues.

Long-Term MAN Growth and Characterization of NPCs

Coat substrate with extracellular matrix Add ECM protein immediately following the final aspiration. Typically we couple fibronectin with poly-L-ornithine and laminin with poly-D-lysine. Matrigel does not require any additional adhesion molecules.

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For fibronectin Fibronectin is provided at 1 mg/ml (0.1% w/v) from Sigma. 6a. Prepare a stock solution of 100 μg/ml in DPBS. Prepare working solution of 10 μg/ml by dilution in DPBS (100 μl stock 1 mg/ml solution in 900 μl DPBS). Store at 4◦ C for up to 6 months from date of receipt; do not freeze. 7a. Completely cover the growth surface of the coverslip or culture vessel with the fibronectin solution and incubate at 37◦ C overnight. Aspirate the ECM solution immediately before use, optionally rinse once with DPBS, and proceed with plating hNPC. See Alternate Protocol 3 for discussion of appropriate plating densities.

For laminin Laminin derived from the basement membrane of Engelbreth-Holm-Swarm mouse sarcoma is provided at 1 mg/ml and laminin from human placental tissue is provided at 0.5 mg/ml (Sigma). 6b. Slowly thaw laminin on wet ice at 2◦ to 8◦ C to avoid gelling. Prepare a 20 μg/ml working solution by dilution in DPBS (20 μl of 1 mg/ml stock murine laminin or 40 μl of 0.5 mg/ml human laminin Per 1 ml DPBS). Store up to 3 days at 4◦ C from date of receipt. Human laminin is used for human cells and murine laminin is used for murine cells. The murine form is much cheaper and both laminins work well, but when using human cells, the authors recommend avoiding mouse proteins. In addition, the murine laminin is from a sarcoma and probably has some minor contaminants in it.

7b. Completely cover the growth surface of the coverslip or culture vessel with the laminin solution and incubate at 37◦ C overnight. Aspirate the ECM solution immediately before use, optionally rinse once with DPBS, and proceed with plating hNPC. Alternatively, aspirate ECM, incubate at 37◦ C overnight, and proceed with plating cells. Laminin and fibronectin solutions may be reused once immediately following coating procedure. See Alternate Protocol 3 for discussion of appropriate plating densities.

For Matrigel Matrigel, growth factor reduced, is provided in 10-ml aliquots (BD Bioscience). 6c. Slowly thaw Matrigel bottle at 4◦ C overnight. Add 10 ml cold Neurobasal medium with a precooled pipet, mix well, aliquot 1 ml per prechilled 15-ml centrifuge tube, and store at −20◦ C. To prepare working solution, slowly thaw 1-ml Matrigel aliquot at 4◦ C for 2 to 4 hr. Add 14 ml chilled Neurobasal medium with a chilled pipet (1:30 final dilution) on ice. Matrigel is extremely temperature sensitive and will prematurely gel if not prepared correctly.

7c. Transfer an appropriate amount of the diluted Matrigel to cover the entire growth surface of the coverslip or tissue culture vessel. Incubate overnight in a Ziploc bag (to prevent evaporation) at 4◦ C. The following day, aspirate Matrigel and immediately seed hNPC at desired concentration. Increasing the concentration of ECM will partially enhance adhesion. Alternatively. precoated ECM coverslips may be purchased from BD Bioscience.

Somatic Stem Cells

See Alternate Protocol 3 for discussion of appropriate plating densities.

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BASIC PROTOCOL 2

ESTABLISHING CLONAL hNPC SUBPOPULATIONS Clonal analysis following mitogen selection and throughout the expansion of hNPC is critical for establishing the cardinal criterion of stemness in newly derived hNPC lines (Vescovi and Snyder, 1999; Gritti et al., 2008). While single-cell cloning provides a method for defining stemness, it does imply that the newly derived subpopulation of hNPC will remain clonally undifferentiated throughout time. In fact, it should be assumed that asymmetric division and subsequent differentiation will occur, resulting in a heterogeneous mixture of hNPC and their differentiated counterparts. For this reason, it is imperative to continually reclone any newly derived hNPC line into smaller subpopulations that can then be functionally tested for multipotency in vitro and in vivo. Achieving clonality of hNPC can be attained by two main methods, limited dilution or flow cytometry. The overall efficiency of either process is extremely low (<5%) for hNPC, due to the extremely slow division rate and requirement of trophic support. In our experience, it has been extremely difficult to amass large quantities of clonal hNPC. Although the first few cell divisions can be enhanced by the addition of CM, medium components eventually break down and salts may begin to come out of solution as time goes on. In addition, it can be extremely arduous to change such a small amount of medium without losing the precious cell cluster. For these reasons, clonally derived hNPC that pass both in vitro and in vivo differentiation assays are extremely valuable and should be meticulously expanded for future use.

Materials Human NPC (Basic Protocol 1 or Support Protocol 1) NB-B-27 complete medium (see recipe) Conditioned medium (CM; Support Protocol 4) Leukemia inhibitor factor (LIF; Millipore, cat. no. LIF1010) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) Normocin (InvivoGEN, cat. no. ant-nr-1) Complete Hibernate-E medium (see recipe) 15-ml conical tubes Fluorescence activated cell sorting (FACS) machine (Robinson et al., 2009) 96-well plates Additional reagents and equipment for dissociating hNPC (Basic Protocol 1 or Alternate Protocol 1) and flow cytometry (Robinson et al., 2009) To clone by limiting dilution 1a. Dissociate hNPC as described in Basic Protocol 1 or Alternate Protocol 1 as applicable. 2a. Plate hNPC by serial dilution (to obtain a calculated concentration of 1 cell/well) into a 96-well plate with 300 μl/well of medium consisting of 50% (v/v) fresh NB-B-27 complete medium/2 μl/ml Normocin/50% (v/v) CM/10 ng/ml LIF/20 ng/ml bFGF. Place in a 37◦ C, 5% CO2 humidified incubator. Cloning can be a very tedious and time-consuming procedure, especially when dealing with slowly dividing hNPC; therefore the basal medium is supplemented with 50% CM for additional autocrine/paracrine support.

Long-Term MAN Growth and Characterization of NPCs

To clone by FACS 1b. Dissociate hNPC as described in Basic Protocol 1 or Alternate Protocol 1, as applicable, and resuspend in 3 ml complete Hibernate-E medium. Hibernate-E is a basal medium that supports short-term hNPC maintenance at ambient carbon dioxide levels (U.S. Patent 6,180,404).

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2b. Sort hNPC by forward and side scatter (Kim and Morshead, 2003; see Robinson et al., 2009, for basic procedures in cell sorting) to create a size gradient at 1 cell/well into a 96-well plate with 300 μl/well of 50% (v/v) fresh NB-B-27 complete medium/50% (v/v) CM/10 ng/ml LIF/20 ng/ml bFGF. Place in a 37◦ C, 5% CO2 humidified incubator. Additionally, sort hNPC using FACS for fluorescently labeled hNPC preparations. hNPC are extremely fragile and do not tolerate mechanical stress well. If possible, the expulsion flow speed should be lowered to increase cloning efficiency.

3. At a time point 6 to 12 hr later, inspect by phase-contrast microscopy (or fluorescence if cells carry fluorescent reporter) to confirm that only one cell resides in each well. Exclude wells containing cellular debris, dead cells, or more than one cell immediately. Within 12 to 24 hr, hNPC will re-equilibrate and appear rounded and phase-bright.

4. Expand clones, supplementing growth factors and heparin every 3 days until small clonal populations expand. Be very careful not to remove or agitate the newly formed cloned cell cluster. Assume that all of the heparin needs to be replaced after 6 to 7 days and that the growth factors and Normocin (2 μl/ml) need to be replaced every 2 to 3 days. At first, growth is very slow, 3 to 4 days per division, with 30% to 50% of total cells dividing at any given time. Generally, it takes months to expand a clone into a 12- or 6-well dish, followed by a month or two to get up to 25-cm2 , then 75-cm2 flasks.

LABELING hNPC PRE-TRANSPLANTATION In order to identify transplanted donor cells, hNPC must be pre-labeled chemically or genetically with a definitive nontransferable marker. One can also utilize human-specific antibodies, such as huNuc, after transplantation to recognize donor-derived cells. For proper validation, hNPC should be identified with at least two of these markers and preferably three to ensure that results are not simply false-positive artifacts. We generally prefer colocalization of BrdU and a reporter gene with at least one human-specific epitope to locate successful donor grafts. Upon implantation into the mammalian CNS, hNPC may undergo one to three rounds of division before becoming post-mitotic; therefore, non-integrating labels will not become too diluted for later detection. Theoretically, thymidine analogs, iron particles, and lipophilic dyes become diluted by a factor of 1/2 for every symmetrical cell division; therefore, these markers can become diluted below standard detection levels within five to six cell divisions. Careful selection of labeling method should be based on the specific assay of interest.

Labeling hNPC with BrdU Human NPC that have not been genetically labeled are preincubated with the Sphase, DNA-intercalating thymidine analog, bromodeoxyuridine (BrdU) for proper postmortem graft identification. Both monoclonal (Gratzner, 1982) and polyclonal antibodies have been raised to detect BrdU using immunofluorescence and multiphoton confocal microscopy for graft analysis. As a result, BrdU was reinforced as a popular prelabeling technique for grafting proliferative cellular substrates. Although BrdU only labels at best 42% to 50% (neurospheres) to 74% to 82% (MAN cultures; Fig. 2D.3.5) of donor hNPC nuclei (in our experience), presents a variety of false-positive artifacts (Rakic, 2002a,b; Burns et al., 2006), and can be highly toxic when administered for extended periods of time (Caldwell et al., 2005), it has remained one of the most highly used pre-transplantation labeling methods to verify donor cell origin (Dolbeare, 1995, 1996; Carbajo et al., 1995; Carbajo-Perez et al., 1995).

BASIC PROTOCOL 3

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Figure 2D.3.5 hNSC MAN cultures incorporate BrdU. Human HSC (HFB-2050) grown as multilayer adherent networks express nestin (A,D) extensively and readily incorporate BrdU (B,E), indicating that they remain in a highly proliferative, immature state throughout MAN culture. Of interest, cells within the clusters appear to proliferate preferentially in comparison to their peripheral counterparts (C,F), indicating a possible niche component within each adherent cluster.

Materials NB-B-27 complete medium (see recipe) Normocin (InvivoGEN, cat. no. ant-nr-1) Leukemia inhibitor factor (LIF; Millipore, cat. no. LIF1010) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) 5-bromo-2 -deoxyuridine (BrdU; Sigma, cat. no. 59-14-3; see recipe) 25-cm2 tissue culture flasks 1. To efficiently label cells, dissociate hNPC 48 to 72 hr prior to transplantation and replate as described in Basic Protocol 1 or Alternate Protocol 1, as applicable, using fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, 20 ng/ml bFGF, and 10 to 20 μM BrdU (added from 1000× stock). BrdU is highly toxic to low-density hNPC cultures; therefore, we recommend plating cells at no less than 2 × 106 cells/25 cm2 . CAUTION: BrdU acts by incorporation in the place of thymidine during DNA synthesis, and thus may cause birth defects or heritable genetic effects. Be extremely careful when handing BrdU. Long-Term MAN Growth and Characterization of NPCs

2. Allow cultures to equilibrate and return to homeostasis in a 37◦ C, 5% CO2 humidified incubator. Low-density cultures fail to equilibrate properly and display extremely slow division rates, impeding efficient labeling during S-phase.

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3. Every 24 to 36 hr, replenish the medium by adding fresh 1000× BrdU to 1× final, LIF to 10 ng/ml final, and bFGF to 20 ng/ml final. Lightly triturate. Gentle trituration of cells is absolutely essential to reduce merging of cell clusters and formation of premature aggregates before transplantation. Cells are extremely fragile when incubating in BrdU; therefore, slow trituration is recommended to avoid shearing.

4. After a total of 48 to 72 hr, prepare hNPC for transplantation as described (see Basic Protocols 6 and 7). In our hands, 40% to 50% (neurospheres) and 70% to 80% (MAN) of the total hNPC population will be labeled after 48 to 72 hr. It should be noted that longer incubation times are notorious for introducing false positives into nondividing cells. Furthermore, hNPC do not proliferate well past 3 to 4 days in media that include BrdU (D.R. Wakeman, unpub. observ.), indicating a time threshold for toxicity.

Lentiviral Infection of hNPC Traditional methods for prelabeling NPC prior to transplantation require harsh DNAintercalating thymidine analogs such as BrdU and CldU, which have been shown to result in great underestimation of overall engraftment success and create a variety of false positives when administered for extensive periods of time. Generation of independently labeled fluorescent reporters, animals, and cell lines eliminates the need for these toxic compounds while increasing both cell viability and engraftment efficiency (Shimomura et al., 1962; Chalfie et al., 1994, 1995; Ward et al., 1998; Zhang et al., 2002; Tsien, 2003; Vintersten et al., 2004; Shaner et al., 2005, 2008; Shimomura, 2005; Giepmans et al., 2006). In addition, fluorescently labeled donor cells can be easily identified among their host counterparts, allowing for enhanced visualization of axonal and dendritic processes. These optical properties also allow us to perform classic electrophysiological assays to test the synaptic potential of differentiated cells and access the overall multi-potentiality of each subline.

BASIC PROTOCOL 4

Lentiviral infection has proven a reliable way to stably express genes of interest (∼8 kb) into slowly dividing hNPC with little to no long-term effects on behavior or morphological phenotype (Consiglio et al., 2004; Capowski et al., 2007; also see UNIT 2D.2). We have used the following protocols to generate hNPC engineered to constitutively express secondand third-generation lentiviruses carrying either a cytosolic CAG-eGFP (kind gift of Mark H. Tuszynski) or PGK-mCherry (kind gift of Mark Mercola) fluorescent marker protein for greater than 5 months with 75% mCherry and 85% eGFP (aggregate) and 89% mCherry and 98% eGFP (MAN) efficiency for aggregate and MAN cultures respectively. All procedures involving live infection-competent lentivirus are performed in a Level 2 or better biosafety hood in accordance with your institution’s specific safety standards. We recommend full disposable safety coat, sleeves, glasses, and double nitrile and latex gloves for adequate personal protection. Any materials (pipets, tips, flasks, conical tubes) that come into contact with virus should be properly sanitized by soaking for at least 20 min in 10% to 20% (v/v) bleach, followed by 15 min in 70% ethanol, and properly disposed of according to safety regulations. Medium is prepared fresh prior to infection. Conditioned medium can be utilized, but metabolic components may influence the overall efficiency in cell lines (D.R. Wakeman, unpub. observ.) and subsequent gene expression (McCarthy et al., 1995). Aggregate cultures can be infected as either single cells at high density or as smallsize clusters (8 to 16 cells/cluster) to medium-size clusters (16 to 32 cells/cluster) (Fig. 2D.3.6). Utilizing cellular aggregates has the added benefit of essential cell-cell contacts, whereas single cells must be plated at high density to induce cell-cell contacts quickly following infection. In addition, larger aggregates contain internal cells that may

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Figure 2D.3.6 Aggregate hNSC cultures express lentiviral eGFP. Human NSC (HFB-2050) were dissociated into single cells at medium density (1 × 106 cells/25 cm2 ) and cultured as lightly adherent aggregates (with trituration every 12 hr) for 48 hr before transfection with a CAG-eGFP lentivirus. 72 hr after exposure (5 days in vitro), free-floating spherical aggregates readily expressed the transgene (A-C). Inset in (C), a close-up of the aggregate in the middle of the frame. When plated onto poly-D-lysine-coated tissue culture–treated flasks, hNPC-eGFP aggregates rapidly attached and flattened, confirming 80% to 90% efficiency, and sustained eGFP expression in vitro (D,E). Green = live eGFP expression.

not be exposed to the viral particles, resulting in decreased infection efficiency. Although most of these cells are migratory within each cluster and eventually become labeled with increased incubation times, we recommend infecting hNPC as either small cellular aggregates or high-density single cells (3–4 × 104 cells/cm2 ), so that all cells are accessible to viral particles for optimal infection efficiency.

Materials

Long-Term MAN Growth and Characterization of NPCs

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Human NPC, single cells or lightly adherent aggregates (Basic Protocol 1 or Alternate Protocol 1) NB-B-27 complete medium (see recipe) Normocin (InvivoGEN, cat. no. ant-nr-1) Leukemia inhibitor factor (LIF; Millipore, cat. no. LIF1010) Basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) 10 mg/ml polybrene (Chemicon) Lentivirus (see protocol introduction for information) Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) Current Protocols in Stem Cell Biology

25-cm2 tissue culture flasks 15-ml conical tubes 1. Dissociate hNPC as described in Basic Protocol 1 or Alternate Protocol 1, as applicable, immediately prior to infection, or utilize small clusters (dissociated 48 to 72 hr before infection, similar to cryopreservation or transplantation). 2. Replate hNPC in 6 ml fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, 20 ng/ml bFGF, and 6 μl of 10 mg/ml polybrene in a 25-cm2 flask. Polybrene enhances the infection efficiency but can be omitted if desired.

3. Carefully add lentivirus at a concentration of 100 ng p24 particles for every 1 × 106 cells. CAUTION: Properly sanitize any virus-exposed waste with bleach and ethanol.

4. Incubate in a 37◦ C, 5% CO2 humidified incubator for 12 to 48 hr. Longer incubation times will result in higher efficiency rates and expression levels, but at the cost of multiple integration sites.

5. To remove any remaining live virus, transfer hNPC to a 15-ml conical tube, rinse the flask once with 10 ml DPBS, combine with the cells in the tube, and allow the cells to settle by gravity. Aspirate the supernatant and repeat 10-ml DPBS rinse three times. Alternatively, centrifuge contents for 4 min at 400 × g in a lentivirus-approved containment centrifuge.

6. After final rinse, add 8 ml fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, and 20 ng/ml bFGF. Incubate in a at 37◦ C, 5% CO2 humidified incubator. 7. Repeat DPBS rinse every 24 hr for the first 7 days to remove any residual viral particles. Excessive trituration and centrifugation can be detrimental to hNPC survival during this crucial time period. We recommend replating single cells as high-density aggregate or MAN cultures to enhance paracrine signaling required for enhanced expansion. Extensive expression of both transgenes is seen after 48 to 72 hr at an efficiency of 82% to 85% for eGFP and 75% to 79% for mCherry single cells and small clusters, respectively. Larger aggregates are much more variable and range between 45% to 85% in labeling efficiency, in the authors’ experience.

Lentiviral Infection of Multilayer Adherent Network (MAN) The MAN assay allows for temporary modification and optimization during lentivirus infections, due to the ease of changing medium; therefore, we reduce the volumes of culture medium to 5 ml fresh hNPC medium in each 25-cm2 flask to concentrate virus and decrease waste. We have applied the following procedure to new and mature (60% to 80% confluent) MAN hNPC cultures (3–5 × 106 cells/flask), infecting with lentivirus containing either a PKC or CAG promoter–driven cytosolic eGFP (Matz et al., 2002; Tao et al., 2007) or mCherry (Merzlyak et al., 2007) fluorescent protein (Fig. 2D.3.7).

ALTERNATE PROTOCOL 4

Additional Materials (also see Basic Protocol 4) 60% to 80% confluent (2 to 3 weeks in vitro) MAN hNPC culture in 25-cm2 flask (Basic Protocol 1) or in transmembrane basket 6-well tissue culture plates Hanging Basket Cell Culture Insert, 1.0 μm (Millipore, cat. no. PIRP30R48) 25-cm2 tissue culture flasks Current Protocols in Stem Cell Biology

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Figure 2D.3.7 MAN cultures efficiently express lentiviral eGFP. Human neural stem cells (HFB-2050) were dissociated into single cells and exposed to a CAG-eGFP lentivirus (as described in text) upon replating (2.5 × 106 cells/25-cm2 flask). hNSC were allowed to attach without agitation to induce formation of multilayer adherent networks (MAN). (A,B) 48 hr post-exposure, hNSC formed small, evenly spaced, adherent aggregate clusters and express the eGFP transgene. (C-H) By 72 hr, transgene expression markedly increased to nearly 90% of hNSC constitutively expressing eGFP throughout the cytoplasm. In addition, adherent clusters continued to proliferate and spread, making initial contacts with neighboring colonies (arrow). (I-L) Over the next week, adherent hNSC aggregates rapidly proliferated and actively migrated between adherent three-dimensional aggregates, creating the initial foundation for the multilayer adherent network. (I-J) Many peripheral anchor cells displayed long protruding feet resembling growth cones that sampled the local microenvironment and rapidly reorganized in response to local guidance cues (arrow). In addition, some clusters detached and continued to proliferate as floating suspension aggregates (arrowhead). (K-N) After 10 days, multilayer adherent clusters began to coalesce, established the classic honeycomb architecture, and actively exchanged migratory proliferative cells between colonies. Individual colonies became unrecognizable as the meandering protrusions of neighboring clusters (arrows) met and rapidly joined into one two-way hNSC highway. (O, P) At 15 days, MAN cultures continued to proliferate and expand across the culturing surface, covering nearly 40% to 60% of the flask by 20 days (Q-S). After 30 days, hNSC (HFB-2050eGFP) assumed mature form (T-V), covering nearly 70% of the culturing surface and were ready to be dissociated. MAN cultures can be maintained for up to 6 weeks; however, overcrowding generally leads to a reduced proliferation rate, and should be avoided if possible. Green = live eGFP expression.

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1. Replace 100% of the medium in 60% to 80% confluent MAN hNPC culture (in 25cm2 flask) with 5 ml fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, 20 ng/ml bFGF, and 5 μl of 10 mg/ml polybrene. Polybrene enhances the infection efficiency but can be omitted if desired.

2. Carefully add the lentivirus at a concentration of 100 ng p24 particles/1 × 106 cells. CAUTION: Properly sanitize any virus-exposed waste with bleach and ethanol. For a 60% to 80% confluent culture, there will be 3–4 × 106 cells per 25-cm2 flask, or 1–2 × 106 cells/ hanging basket cell culture insert.

3. Incubate 12 to 48 hr in a humidified 37◦ C, 5% CO2 incubator. Longer incubation times will result in higher efficiency rates and expression levels but at the cost of multiple integration sites.

4. Remove any remaining virus by rinsing five times with 5 ml DPBS. One of the greatest benefits of utilizing the MAN assay is the ease of changing medium and rinsing live virus from the culture without extensive manipulation or harsh mechanical stress. Using PBS without salts often leads to detachment of cells from the flask; therefore MAN cultures are always rinsed with DPBS containing both Mg2+ and Ca2+ .

5. Add 8 ml fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, and 20 ng/ml bFGF. Incubate in a humidified 37◦ C, 5% CO2 incubator. 6. Repeat steps 4 and 5 every 12 hr for the first 7 days to remove any residual viral particles. Theoretically, all live virus will be removed with the first few rinses; however, we err on the side of safety to ensure adequate removal of all viral particles over time. In fact, we treat lentiviral-infected cultures as if they contain live virus until the cultures have been fully dissociated and passaged at least three times.

7. After 1 to 2 weeks in culture, dissociate (1:3) and expand the newly labeled hNPC population. Extensive expression of both transgenes was seen after 48 to 72 hr at an efficiency of 98% eGFP and 89% mCherry. After three to five additional population doublings, fluorescently labeled hNPC populations are pooled together and sorted by FACS into three polyclonal populations based on relative fluorescence intensity, labeled low-, medium-, and highintensity expression. These cells are then expanded in 25-cm2 or 75-cm2 tissue-culturetreated flasks (Falcon) as MAN or lightly adherent aggregate cultures and frozen for future expansion. Polyclonal subpopulations can then be individually subcloned (Basic Protocol 2) and transplanted intraventricularly into neonatal (P0) mice to assay both hNPC migration and differentiation capacity.

8. As an example, perform MMS lentiviral infection (see Alternate Protocol 2) of hNSC (HFB-2050) as follows. The human NPC line HFB-2050 was engineered to constitutively express either a cytosolic eGFP or mCherry fluorescent marker protein by the following methods. After 20 days of culture, hNPC MMS-multilayer adherent networks were infected with lentivirus containing either a cytosolic eGFP or mCherry protein driven by the CAG or PKC promoters respectively (Fig. 2D.3.8).

a. Because the MMS allows for temporary modification and optimization during lentivirus infections due to the basket’s inherent mobility, reduce the volume of culture medium to 1.5 ml fresh NB-B-27 complete medium containing 2 μl/ml Normocin, 10 ng/ml LIF, 20 ng/ml bFGF in each well of a 6-well dish, and insert a transmembrane basket with attached hNSC into each well. For a 60% to 80% confluent culture, there will be 1–2 × 106 cells per hanging basket cell culture insert.

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Figure 2D.3.8 MAN and MMS mCherry lentivirus expression. Human NSC (HFB-2050) grown as multilayer adherent aggregate (MAN) cultures maintain expression of the PGK-mCherry transgene 10 days post transfection (A-D). In addition, the MAN culturing technique can be adapted to a semi-porous membrane culturing surface and grown using the MMS technique, highly resembling MAN cultures in phenotype and composition. MMS cultures are then easily transfected with viral vectors (as described in the text) to produce fluorescent reporter cultures that are easy to manipulate (E-G).

b. To each basket insert, add 10 mg/ml polybrene and lentivirus at a concentration of 100 ng p24 particles/1 × 106 cells. Incubate in humidified 37◦ C, 5% CO2 incubator. CAUTION: Properly sanitize any virus-exposed waste with bleach and ethanol.

c. At a time point 48 hr later, aspirate viral waste and wash hNPC five times for 30 sec each in 5 ml DPBS. Transfer the basket insert to a new 6-well plate and return to standard culture medium volumes (see Alternate Protocol 2). This procedure is repeated every day for 7 days to remove any residual viral particles. Extensive expression of both transgenes was seen in HFB-2050 after 72 hr at an efficiency of 98% eGFP and 89% mCherry (quantified by FACS).

d. After 7 days, dissociate hNPC networks into single-cell suspension by adding 1.5 ml per well of Accutase and seed one well into one 25-cm2 flask with 8 ml NB-B-27 medium containing 2 μl/ml Normocin, 10 ng/ml LIF, and 20 ng/ml bFGF per flask, to induce MAN or lightly adherent aggregate cultures.

Long-Term MAN Growth and Characterization of NPCs

In the case of HFB-2050, each well was dissociated into a 25-cm2 flask, cultured, and expanded as a webbed MAN for 20 days before passaging again into one 75-cm2 . After three total passages (90 days), all hNSCs were pooled together and sorted by FACS into three polyclonal populations based on relative fluorescence intensity, labeled low-, medium-, and high-intensity expression. These cells were then expanded in 25-cm2 or 75-cm2 tissue culture–treated flasks (Falcon) in modified adherent networks or as suspension aggregates, frozen for future expansion, or cloned and transplanted into (P0) neonatal mice intraventricularly (Basic Protocol 8) to assay both migration and differentiation capacity.

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Labeling hNPC with Super-Paramagnetic Iron Oxide (SPIO) To follow survival and migration of transplanted donor hNPC in vivo, it has become increasingly important to develop noninvasive techniques (Manganas et al., 2007; Gilad et al., 2008; Ruiz-Cabello et al., 2008) for monitoring and imaging engrafted donor cells (Rogers et al., 2006; Slotkin et al., 2007; Schroeder, 2008; Sumner et al., 2007). Common post-mortem immunohistological staining techniques do not afford the opportunity to trace migration within the same animal over time; however, hNPC transfected with super-paramagnetic iron oxide (SPIO) particles have been used with MRI in rodents to successfully track migration of stem cells after engraftment in the normal and diseased animal (Lewin et al., 2000; Bulte et al., 2002; Frank et al., 2003; Hinds et al., 2003; Arbab et al., 2003a,b, 2004a,b; Jendelova et al., 2004; Miyoshi et al., 2005; Magnitsky et al., 2005; Sykova and Jendelova, 2005, 2006, 2007a,b; Lepore et al., 2006a,b; Shapiro et al., 2006; Guzman et al., 2007, 2008; Politi et al., 2007; Walczak and Bulte, 2007; Neri et al., 2008; Walczak et al., 2008). Successfully labeled cells can be detected and followed weeks to months after implantation as they migrate contralaterally through predominantly white matter to sites of pathological insult. In addition, MRI findings can then be verified post-mortem utilizing immunohistochemistry by costaining for human specific markers and iron particles (Prussian blue or dextran staining) utilizing the MRIguided coordinates.

BASIC PROTOCOL 5

Additional Materials (also see Basic Protocol 4) Human NPC (Basic Protocol 1 or Alternate Protocol 1) Feridex (Bayer Healthcare Pharmaceuticals, cat. no. NDC-59338-7035-5) Protamine sulfate injection, USP, 50 mg/5 ml (Bayer Healthcare Pharmaceuticals, cat no. NDC-63323-229-05) 25-cm2 tissue culture flasks 15-ml conical tubes Centrifuge 1. Dissociate hNPC into single cells as described in Basic Protocol 1 or Alternate Protocol 1, as applicable. Replate 2–3 × 106 cells in a 25-cm2 flask containing 5 ml of 25% CM/75% fresh NB-B-27 complete medium /10 ng/ml LIF/20 ng/ml bFGF/2 μl/ml Normocin. Incubate for 6 hr at in a 37◦ C, 5% CO2 humidified incubator. Alternatively, skip the incubation and proceed directly to step 2. We prefer to allow freshly dissociated hNPC to recover for 6 to 12 hr after dissociation to increase viability before SPIO incubation.

2. Prepare 2× SPIO mixture, 45 to 60 min before labeling, by adding 10 to 20 μg/ml Feridex and 5 μg/ml protamine sulfate to 5 ml fresh NB-B-27 growth medium containing 10 ng/ml LIF and 20 ng/ml bFGF. Incubate in a 37◦ C water bath for 45 to 60 min with mixing. We have had the best short-term in vitro results utilizing a final working concentration of 10 μg/ml Feridex.

3. Add 5 ml of 2× SPIO solution to the medium over the plated cells from step 1 and triturate gently for a final working concentration of 5 to 10 μg/ml Feridex/2.5 μg/ml protamine sulfate. Incubate for 20 to 24 hr in a 37◦ C, 5% CO2 humidified incubator to allow iron particles to enter the cell by endosomal transport. Increasing the Feridex concentration will enhance the overall signal but can result in intracellular clumping and aggregation of iron particles that may be detrimental to the long-term viability, stability, and behavioral phenotype of the cell. Somatic Stem Cells

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4. After 20 to 24 hr, gently triturate hNPC and transfer the contents to a 15-ml conical tube. Rinse the flask once with 4 ml DPBS and add to the conical tube. Centrifuge for 3 to 4 min at 400 × g, room temperature. Increasing the incubation time will enhance the number of iron particles that enter the cell by only a small percentage and possibly at the cost of cellular differentiation or decreased post-incubation survival. In addition, the total number of particles per cell will vary within the individual culture. 24 hr is sufficient for nearly 95% labeling efficiency.

5. Vacuum aspirate the supernatant and rinse by resuspending in 8 ml DPBS. Centrifuge for 3 to 4 min at 400 × g, room temperature. Repeat DPBS rinse twice. 6. Resuspend the SPIO-labeled hNPC pellet in 8 ml fresh prewarmed NB-B-27 complete medium/10 ng/ml LIF/20 ng/ml bFGF/2 μl/ml Normocin. Replate in a 25-cm2 tissue culture flask and incubate in a 37◦ C, 5% CO2 humidified incubator for 6 to 12 hr, or proceed immediately to transplantation procedures (see Basic Protocols 6 and 7). Alternatively, replate SPIO-labeled hNPC onto Matrigel-coated 24-well plates (Support Protocol 5) for in vitro characterization and differentiation assays. Recommended plating densities are 50,000 cells/well for a 24-well plate, 100,000 cells/well for a 12-well plate, or 250,000 cells/well for a 6-well plate. The hNPC may also be labeled 24 hr after plating onto Matrigel by simply adding the SPIO mixture to the newly established adherent monolayer culture. SUPPORT PROTOCOL 6

Perls Prussian Blue Staining (for Hemosiderin) Detection of intracellular SPIO particles within hNPC can be achieved by a chemical reaction that converts ferric iron to an insoluble blue compound called Prussian blue or Berlin blue (Fig. 2D.3.9). Briefly, addition of HCl forms ferric chloride, which is then converted to the insoluble ferric ferrocyanide (Prussian blue) with potassium ferrocyanide. The color intensity CANNOT be used quantitatively to assess total SPIO input; however, it is commonly used as a qualitative measure for visual comparison. Alternatively, immunostaining for iron particles can be achieved using anti-dextran antibodies, but with a decreased labeling efficiency as compared to Prussian blue nuclear staining (Frank et al., 2007). The chemical reaction is: 4 FeCl3 + 3 K4 Fe(CN)6 → Fe4 [Fe(CN)6 ]3 + 12 KCl.

Materials Potassium ferrocyanide HCl, concentrated Sample: SPIO-labeled hNPC (Basic Protocol 5) or cardiac-perfused (Basic Protocol 8) tissue sections from transplanted animals (Basic Protocol 7 or 8) Fixative: e.g., 4% paraformaldehyde (PFA; see recipe) or equivalent Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) Nuclear counterstain, e.g., neutral red, DAPI, propidium iodide 1. Prepare solutions 1 and 2 and the working solution fresh on the day of staining in iron-free containers.

Long-Term MAN Growth and Characterization of NPCs

Solution 1: 2 g potassium ferrocyanide 100 ml distilled water Solution 2: 2 ml concentrated HCl 100 ml distilled water Working solution: One part solution 1 + one part solution 2.

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Figure 2D.3.9 hNPC readily engulf SPIO particles in vitro. Human NSC (HFB-2050) were plated onto Matrigel with SPIO particles for 20 hr and processed with Perl’s Prussian blue to detect labeled cells. Most hNSC readily took in the Feridex particles (blue particles) (A,B), whereas unlabeled control cells (C,D) were completely devoid of blue staining.

2. Fix hNPC in 4% PFA or another suitable iron-free solution for 20 min prior to staining to avoid false-positives (tissue sections from transplanted animals will have been previously cardiac perfused, as in Basic Protocol 8). Do not rinse sections in tap water, as rust in the water or pipes may alter results. 3. Add working solution to sample until fully submerged and incubate at room temperature for 15 min. Iron particles will develop into blue spots with labeled cells as the reaction proceeds. Use a positive control sample to determine optimal incubation and inactivation time.

4. Aspirate solution and rinse five times with DPBS to remove residual stain. 5. Stain with neutral red, DAPI, propidium iodide, or any other appropriate nuclear counterstain for 2 to 5 min. Rinse thoroughly with DPBS to remove residual stain. Ferric iron (SPIO) will be visible as intracellular Prussian blue precipitates and nuclei as red or blue depending on counterstain chosen.

PREPARING hNPC FOR TRANSPLANTATION Human NPC injections are performed with cells in a uniformly undifferentiated state, displaying log-phase growth, and able to incorporate BrdU during DNA synthesis. To enhance synchronization, undifferentiated hNPC are dissociated 2 days prior to

BASIC PROTOCOL 6 Somatic Stem Cells

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transplantation and disaggregated into a single-cell suspension immediately prior to surgery. Maintaining hNPC in vitro for a longer period of time prior to injection promotes clumping due to spontaneous elaboration of extracellular matrix (ECM) or subsequent premature spontaneous cellular differentiation, both of which may deter efficient engraftment. Careful dissociation of hNPC is essential for sustained viability and longterm post-transplantation survival. Many global neurodegenerative disorders affect multiple CNS cell types; therefore, undifferentiated hNPC provide the plasticity needed for the host microenvironment to naturally direct differentiation into multiple phenotypes that may be necessary to ameliorate and restore the host cytoarchitectural milieu. On the other hand, focal CNS disorders may benefit more from lineage-directed pre-differentiation strategies, as a priori ex vivo manipulation of hNPC along specific neuronal fate pathways may be more suitable for alleviating lineage-specific CNS deficits. These cells are presumably lineage-restricted and committed, effectively eliminating nonspecific, regionally inappropriate differentiation that could lead to potentially detrimental off-target side effects. In contrast to their undifferentiated hNPC counterparts, pre-differentiated precursors and mature post-mitotic neurons to do not have the same capacity for long-distance migration or the multipotency beneficial to global neurodegenerative diseases (Le Belle et al., 2004). The decision to implant a semi-homogenous population of undifferentiated hNPC or lineage-defined progenitors is ultimately determined by the specific cellular properties that best suit the experimental paradigm (Svendsen et al., 1997a; Armstrong et al., 2000; Teng et al., 2002; Pluchino et al., 2003; Burnstein et al., 2004; Kelly et al., 2004; McBride et al., 2004; Yasuhara et al., 2006; Lee, H.J. et al., 2007; Lee, J.P. et al., 2007; Tarasenko et al., 2007). Both strategies have their merits, allowing the natural host tissue to effectively direct maturation and terminal differentiation in accordance with local signaling cues (Fricker et al., 1999).

Materials Human NPC (hNPC): small to medium aggregate clusters (Basic Protocol 1 or Alternate Protocol 1) Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) 0.4% (v/v) trypan blue Hemacytometer (UNIT 1C.3) Additional reagents and equipment for counting viable cells using a hemacytometer and trypan blue (UNIT 1C.3) 1. Triturate hNPC lightly against the bottom of the flask to detach any cellular aggregates that may have attached, and transfer the contents to a 15-ml conical tube. Centrifuge for 3 to 4 min at 400 × g, room temperature. Utilize small- to medium-size aggregates plated 48 hr before transplantation.

2. Dissociate hNPC by the methods previously described (Basic Protocol 1 or Alternate Protocol 1, as applicable) and resuspend in 10 ml cold DPBS. Dissociation should not take longer than 1 to 3 min to fully break apart small aggregates.

3. Centrifuge the 15-ml conical tube for 4 min at 400 × g, room temperature. Remove supernatant. Repeat steps 2 to 3. 4. With a pipettor and 100-μl pipet tip, carefully resuspend the hNPC pellet in a small volume of cold DPBS (20 to 40 μl), approximately equal to the volume of the cell pellet, and transfer into a 1.5-ml microcentrifuge tube. Long-Term MAN Growth and Characterization of NPCs

Resuspend hNPC in half the volume of DPBS first, then wash the 15-ml tube with the second half volume and transfer all cells into the 1.5-ml microcentrifuge tube. Be very careful not to triturate aggressively, as hNPC will easily shear when resuspended at such high density.

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5. Remove 2 μl of cell solution and transfer to a microcentrifuge tube containing 17 μl DPBS, add 1 μl of 0.4% trypan blue to the tube, and triturate well. 6. Count the cells with a hemacytometer, making sure to factor in the 1:10 dilution (UNIT 1C.3). The purpose of this dilution is to use the fewest cells possible for counting, while maintaining accuracy.

7. Adjust the amount of DPBS in the microcentrifuge tube as needed to achieve the desired cell concentration. We aim for ∼50,000 cells/μl as an optimal concentration for transplantation. Cells should not be suspended at higher than 100,000 cells/μl for long periods of time, as excessive cell death will likely occur, resulting in a sticky DNA precipitate that will easily clog the needle.

8. Immediately following resuspension in DPBS, place the vial of hNPC onto wet ice (4◦ C) and gently flick the tube every minute to deter clumping of cells that may clog the needle during stereotactic injection. Proceed without delay to Basic Protocol 7. Cells should be utilized as quickly as possible to reduce cell death and increase overall viability and engraftment success. For multiple animal studies, it is beneficial to prepare separate biological replicate batches as the procedures continue, preparing enough cells for use no longer than 1 hr post-dissociation.

LOADING AND INJECTION OF hNPC FOR TRANSPLANTATION INTO ST. KITTS AFRICAN GREEN MONKEY

BASIC PROTOCOL 7

The following methods have been refined over a number of years for stereotaxic injection of cells and tissue into the nigrostriatal system of monkeys. There are some variations in the methods which other investigators have used, but this procedure has evolved to work successfully in our hands (Redmond et al., 1986, 1988; Taylor et al., 1995; Sladek et al., 1995, 2008; Bjugstad et al., 2005, 2008; Wakeman et al., 2006; Redmond et al., 2007, 2008). NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Committee for Ethical Animal Care and Use (IACUC) and must conform to government regulations for the care and use of laboratory animals.

Materials Experimental animals: St. Kitts African Green Monkey (MPTP-treated or PBS sham control) Ketamine hydrochloride injection, USP (Ketaset, Fort Dodge Animal Health) Atropine for i.m. or s.c. administration Pentobarbital for i.v. administration Lubrivet (optional; Butler Animal Health; http://www.accessbutler.com/) Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) 70% ethanol Human NPC (hNPC; Basic Protocol 6), labeled according to any of the pretransplantation labeling procedures described in this unit NB-B-27 complete medium (see recipe) without Normocin Phosphate-buffered saline (PBS) 100-μl syringe (Hamilton) for stereotactic injection 22-G, 2-in. needle (Hamilton) for stereotactic injection Single syringe microinjectors for Hamilton syringes (Stoelting)

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Primate or large animal size stereotaxic head holder and bars (David Kopf Instruments) Ear bars (David Kopf Instruments) Animal clippers Endotracheal tube Sterile surgical supplies including: Stereotactic bone drill with 0.5-mm drill burrs Drapes Forceps Hemostats Needle holders Periostial elevators Scalpel with no. 10 surgical blade (Bard–Parker) Self-retaining retractor Towel clips Standard neurosurgical supplies—sutures, bone wax, Gelfoam, sterile NaCl irrigation solution 3-in. sterile gauze EKG machine with monitoring leads Leads for temperature and O2 saturation Additional reagents and equipment for counting viable cells with a hemacytometer and trypan blue (UNIT 1C.3) 1. Mount the syringe into the stereotactic apparatus, calibrate according to ear bar zero, and calculate target sites from atlas or prior studies. For injecting monkeys, we steam-autoclave all of the equipment in the surgical field, including the stereotaxic frame with the carriers. The syringe pumps, syringes, and controllers are sterilized by ethylene oxide gas.

2. Anesthetize monkeys initially with 7 to 15 mg/kg ketamine by i.m. injection and administer 0.02 to 0.05 mg/kg atropine by i.m. or s.c. injection. Shave and prepare the surgical area. Place an intravenous line into the saphenous or other peripheral vein and induce the animal to light anesthesia with 20 to 30 mg/kg pentobarbital for placement of an endotracheal tube to ensure adequate airway and for supplemental ventilation with room air or O2 , if necessary. 3. Place the anesthetized and monitored monkey into the stereotaxic frame with the ear bars (it helps to lubricate and clean the ear canals with Lubrivet) and check the centering. Repeat if necessary. For accuracy of targeting, this is the most critical step of the procedure, if it is performed without MRI targeting.

4. Place EKG monitoring leads and a lead for temperature and O2 saturation. 5. Scrub and prep the already shaved head from the eyebrows to the back of the neck and from ear to ear. Remove the drape which protects the sterile cover over the ear bar on the stereotaxic frame, and then drape the head with sterile drapes and an incise drape so that nothing remains exposed except the sterile ear bar, which remains accessible for the syringe holder/carrier. 6. Recheck the zero coordinates for the drill and for the syringe carriers against a calibrated zero bar. Long-Term MAN Growth and Characterization of NPCs

7. Compare the midline of the skull with the calculated coordinates for the drill. Adjust if necessary and then recalculate all targets for the drill and the carriers if the midline

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was adjusted. After the coordinates have been verified, slowly drill holes, lubricating with sterile DPBS. Replace the drill holder with the needle carrier and repeat steps 6 to 7. 8. Clear the Hamilton syringe by drawing and expelling 60 μl sterile DPBS two to three times. The DPBS rinse removes residual ethylene oxide, helps lubricate the needle, and ensures proper working condition of the syringe

9. To fill the needle, wipe the hNPC suspension tube (optimally containing ∼50,000 cells/μl) thoroughly with 70% ethanol and carefully uncap the tube inside the surgery suite. Gently flick or triturate the hNPC microcentrifuge tube immediately prior to filling the needle.

10. Grasp the microcentrifuge tube with hemostats and carefully move the tube up to the needle from underneath. The bevel of the needle should be located at the middle of the cell suspension.

11. Slowly move the tube vertically to mix cells and begin drawing 10 to 20 μl hNPC into the needle with the controlled-rate syringe pump. Cells should be drawn into the syringe immediately before the injection. Do not touch the sides or bottom of the tube, introduce air bubbles, or bend the needle. Prolonged time within the syringe will result in clumping and subsequent clogging of the needle. Always draw 2 to 3 μl excess cell suspension into the syringe for post-injection analysis.

12. Immediately before insertion of needle, slowly expel 1 μl cell suspension from needle into a clean sterile microcentrifuge tube. Add 1 ml fresh NB-B-27 complete growth medium without Normocin to the microcentrifuge tube. 13. Lower the needle into target area of interest at slow speed (10 to 15 sec) to the proper depth for the given stereotactic location. At the appropriate vertical depth, wait 2 min to allow the brain tissue to adjust to the proper level at the needle tip.

14. Inject the prepared cell suspension using the syringe pump at a maximum constant rate of (1 μl/min). 15. During autoinjection, count viable hNPC in the microcentrifuge tube with a hemacytometer and trypan blue (UNIT 1C.3) to determine viability at time of injection and replate the 1 ml of hNPC into one well of a 24-well dish. Incubate at 37◦ C for 48 to 72 hr and examine under the microscope to determine if any contamination was introduced during the procedure. 16. After the cells are fully expelled from the syringe, allow the needle to remain in place for 2 min before withdrawing. The extra settling time will prevent significant backflow and leakage of hNPC through the needle track during retraction of the needle.

17. Retract needle slowly, at a maximum of 1 μm/min. If another injection is planned, flush out the syringe with sterile PBS. Refill and repeat step 8 above. 18. After the last injection, make sure that any bleeding is controlled, suture animal, and proceed to post-operative care. Somatic Stem Cells

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BASIC PROTOCOL 8

INTRAVENTRICULAR INJECTION OF hNPC INTO NEONATAL MICE Neonatal mice (post-natal day 0 to 3; P0 to P3) are relatively effortless to handle and can easily be injected in the lateral ventricles with hNPC, offering several advantages over adult mice for transplantation. The undeveloped newborn skull is soft and translucent, obviating the need for drilling or cutting into the head while facilitating penetration of the needle or drawn glass micropipet. In addition, hNPC survive, migrate, and integrate within the developing neonatal CNS at a higher efficiency than when transplanted into the mature adult brain. It should be noted that engraftment, migration, synaptic maturation, and development of fully competent neuronal subtypes often take longer with human-derived NPC than with rodent-derived NPC; therefore, we highly recommend the use of immunodeficient rodent models such as the SCID genetic background to increase overall success and eliminate the need for expensive immunosuppressants (Chidgey et al., 2008). Other strains with variations in T, B, and NK cell deficiencies such as BALB/c and Rag2 backgrounds may also be used, but may display some leakiness with age. We prefer the NIHS-beige-nude-xid (NIHBNX) or C.B-17 scid beige (CBSCBG) mice available from Taconic or Harlan. The extra cost for maintenance of these animals is far outweighed by time and resource expenditures involved in continually testing and rederiving new hNPC lines, which could be avoided by eliminating the false negatives involved in using nonimmunopriveleged strains.

Materials Neonatal mice (P0 to P3) of appropriate strain 70% ethanol Dulbecco’s PBS with Ca2+ and Mg2+ (DPBS; Mediatech, cat. no. 21-030-CM) Human NPC (hNPC; Basic Protocol 6; 50,000 cells/μl) in microcentrifuge tube Borosilicate glass (Sutter Instrument Co., cat. no. B100–75-15) Micropipet puller (Sutter Instrument Co., Model P-87) Aspirator tube assemblies for calibrated microcapillary pipets (Sigma Aldrich, cat. no.A5177–5EA) Fiber optic light source for transillumination (Dolan-Jenner Industries) Warm-water glove balloon Additional reagents and equipment for preparing injection micropipet (Lee et al., 2008) and processing mouse brains (Basic Protocol 9) 1. Prepare calibrated drawn borosilicate glass micropipet using borosilicate glass and a micropipet puller (Lee et al., 2008). 2. Anesthetize the neonatal mouse by placing the pup on wet ice for ∼1.5 to 3 min until the animal no longer retains locomotion or responds to gentle toe and tail pinch. Carefully monitor the pup to minimize time on ice, as overexposure to low temperatures can lead to death of newly born mice. Immediately proceed to transplantation

3. Insert a calibrated drawn borosilicate glass micropipet into an aspirator tube assembly. Just prior to drawing up hNPC, rinse the micropipet by drawing up and then expelling 5 μl of 70% ethanol ten times, followed by sterile DPBS ten times, to sterilize and lubricate the glass.

Long-Term MAN Growth and Characterization of NPCs

4. Gently flick or triturate the microcentrifuge tube containing the hNPC suspension immediately prior to filling the needle. Wipe the tube thoroughly with 70% ethanol and carefully uncap the tube.

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5. Slowly move the tube vertically to mix cells and begin drawing 4 to 5 μl hNPC into the micropipet by mouth suction. Do not touch the sides or bottom of the tube, introduce air bubbles, or break the glass needle. Prolonged time within the micropipet will result in clumping, subsequently clogging the needle. Always draw 2 to 3 μl excess cell suspension into the syringe for post-injection analysis.

6. Loosely secure the skull by hand at the cranium and place directly over a non-heatconducting (fiber-optic) light source to visualize the bregma and lateral ventricles by transillumination. Gentle handling of the pup throughout the procedure is important to avoid trauma.

7. Carefully insert glass needle ∼0.5 to 1 mm deep into the head at the midline between bregma and eye and slowly inject 1 to 2 μl hNPC suspension at 5 × 104 cells/μl into the lateral ventricle of either the left or right hemisphere. Correct location and accurate dispersion within the ventricles can be confirmed by addition of an inert dye to the cell suspension. A correctly placed glass needle will deliver cells without resistance by mouth-pressure delivery from a micropipet aspirator tube assembly. We recommend practicing on nonexperimental animals with trypan blue, to become acquainted with the correct pressure and distance necessary for accurate intraventricular injection of experimental animals with hNPC cell suspensions. P0 mice generally will tolerate 3 μl total divided between both ventricles, while the later stages P2 to P3 can tolerate 4 to 5 μl total.

8. Slowly remove the needle and check for leakage through the needle track. Repeat step 7 in the contralateral hemisphere. The entire procedure starting from grasping the pup off of ice to injection should not take longer than 40 to 60 sec per animal to avoid having the anesthetic wear off.

9. Immediately following injection, warm the lower extremities under tepid flowing water, gently pat dry with a cotton swab or piece of gauze, and place on a warm-water glove balloon on top of a heating pad to adequately increase the body temperature of each pup before returning the pup to its mother, to avoid post-operation parental rejection. Monitor pups after transplantation for several hours to ensure adequate post-operational recovery. It is very important to increase core body temperature quickly, for best results. The brain is not an immunopriveleged organ; therefore immunosuppression regimes are recommended to avoid rejection in applicable genetic backgrounds (e.g., wild-type strains).

10. Process brains for differentiation potential at the following time points (see Basic Protocol 9): a. Injection location confirmation (2 to 12 hr). Location and structures of interest, lateral ventricles and luminal cavity: Trypan blue tracer can easily be seen within the lateral ventricles at this time, confirming correct needle position at time of injection. In addition, the overall integrity of the brain and amount of damage incurred by the glass needle track can be assessed to validate technical procedure for optimal hNPC transplants. Cells will begin to engraft in the first 8 to 12 hr following transplantation; however, accurate assessment for cell survival can not be attained this early.

b. Engraftment (12 to 24 hr). Location and structures of interest, lateral ventricular walls, including SVZ and choroid plexus:

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Cells will initially incorporate within the ventricular walls and preferentially accumulate in the SVZ as bulbous nodules where they eventually incorporate into the host SVZ and rostral migratory stream (RMS). Dead cells are often cleared and become trapped within the choroid plexus, the natural “kidney” of the brain that actively filters CSF. In general, there is a give and take between the moderate amount of cell death that occurs during the transplantation process and the subsequent cell divisions that take place post-transplantation.

c. Integration and migration within SVZ-RMS niche (1 to 3 days). Location and structures of interest, SVZ, RMS, olfactory bulbs (OB): Successfully engrafted hNPC incorporate within the developing ventricular walls, forming nodules reminiscent of SVZ protrusions induced by intraventricular growth factor infusion (Kuhn et al., 1997; de Chevigny et al., 2008). The bulbous cluster eventually flattens back out into a normal ventricular wall, and the hNPC integrate within the host neurogenic niche. Within the endogenous SVZ niche, a small population of donor cells will continue to proliferate (Ostenfeld et al., 2000) for at most two to four cell divisions (in our experience) before exiting the cell cycle as they mature and coordinately migrate tangentially and by chain migration through the RMS. The number of cells that die during and following transplantation far outweighs the additional proliferative load; therefore, unchecked tumor-like growth does not occur.

d. Migration, differentiation, and synaptic integration (3 to 14 days). Location and structures of interest, RMS and olfactory bulbs (OB), neuraxis: As hNPC preferentially migrate through the RMS to the OB, they receive signaling and guidance inputs that eventually direct them to defer from tangential or chain migration and turn radially where they continue to migrate and differentiate into synaptically integrated OB interneurons. In addition, many cells will migrate through predominantly white matter and integrate appropriately into the developing striatum and to a lesser extent contribute to the cortex as well (Fig. 2D.3.10). Utilizing non-immunopriveleged animals that do not exhibit pathological deficit often leads to a significant decrease in overall graft survival, integration, and long-term synaptic connectivity. When testing hNPC differentiation potential, we recommend always using immunodeficient mouse models (SCID, BALB/c, or Rag2) to decrease time-consuming false negatives. Assaying donor cell survival after 2 to 3 weeks in vivo in nondiseased animals can often be misleading, as many of the cells are discarded, become quiescently integrated within the ventricular wall and SVZ, or simply undergo apoptosis in response to local microenvironment niche signaling cues. It seems that once the initial process of fetal development has concluded, only a small portion of transplanted hNPC remain quiescently undifferentiated within the neurogenic niches, while the remainder of donor cells continue to migrate, differentiate, and eventually be replaced by the natural host neurogenic process. In non-immunopriveleged animals, we speculate that these donor cells undergo additional selectional pressure as the host immune system develops, and this may eventually deter the long-term survival and maturation of OB interneurons. One way or another, very few transplanted hNPC will be found after 1 to 3 months in vivo in nondiseased, non-immunopriveleged animals; therefore, careful consideration of genetic background and terminal end point is essential for transplantation success.

Long-Term MAN Growth and Characterization of NPCs

Figure 2D.3.10 (at right) Engrafted hNSC survive, mature, and express the eGFP transgene in vivo. Human NSC (HFB-2050) engineered to express eEGP under the CAG promoter were established and transplanted at low density into both lateral ventricles of P0 neonatal mice (as described in the text). Mature eGFP+, donor-derived neurons were found in clusters throughout the forebrain up to 10 weeks post-transplantation (A-H); DAPI (B,F), phase (C,G), GFP (A,D,E,H).

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Figure 2D.3.10

(legend at left)

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BASIC PROTOCOL 9

PROCESSING ENGRAFTED MOUSE BRAINS Animals receiving hNPC transplants should be sacrificed and processed with strict consideration for the specific assays and questions of interest to the investigator. Engrafted brains can be sectioned in the coronal or sagittal plane with either a cryostat (for samples that require low temperature, or to create ultrathin sections) or vibratome (standard immunoassays) and processed for further analysis.

Materials Injected experimental animals (Basic Protocol 8) Anesthetic: Isoflurane or pentobarbital Dulbecco’s PBS without Ca2+ or Mg2+ (CMF-DPBS; Mediatech, cat. no. 21-031-CM) 4% paraformaldehyde (PFA; see recipe), cold 10% and 30% (w/v) sucrose in PBS OCT embedding medium (e.g., Fisher) Dry ice 2-methylbutane (Fisher, cat. no. 03551-4) Sodium azide Low-melt agarose (UltraPure Low Melting Point Agarose; Invitrogen, cat. no. 16520050) Dissection tools: scalpel, fine scissors, forceps, spatula, and pins Perfusion Apparatus: pump and leads Appropriate embedding mold Charged slides (Fisher Superfrost Plus) Cryostat microtome or vibratome 1. At time point of interest, deeply anesthetize animal using inhalant isoflurane or 40 to 85 mg/kg pentobarbital injected i.p., then sacrifice by live cardiac perfusion in cold CMF-DPBS to clear blood, followed by perfusion with fresh cold 4% PFA for fixation. Ensure animal is fully anesthetized as assessed by toe and tail pinch. It is important to clear blood from the vessels within the brain to eliminate background autofluorescence during imaging. Perfusions should always be performed with cold reagents. Ensure that full rigor mortis has set in before completion of perfusion. The tail will curl upon introduction of PFA if the perfusion leads are correctly positioned.

2. Carefully dissect out and remove brain, then submerge in fresh cold 4% PFA for 24 to 48 hr at 4◦ C for deep fixation. Dissect the whole brain, paying careful attention not to sever the cerebellum, spinal cord, or olfactory bulbs. We recommend starting from the hindbrain side and moving rostrally toward the OB. OBs can be safely removed intact and attached to the forebrain by carefully separating the meninges and coaxing the OB from the nasal cavity area with a small rounded flat spatula.

To cryopreserve and cryosection Cryosectioning should be employed for sectioning extremely thin slices, such as when used in electron microscopy or when assays require preservation of tissue at low temperatures. 3a. Soak brains for 24 hr at 4◦ C in CMF-DPBS containing 10% sucrose, followed by 24 hr at 4◦ C in PBS containing 30% sucrose. Long-Term MAN Growth and Characterization of NPCs

4a. Place the brain into an appropriate mold and pour OCT solution into the mold so that the entire brain is submerged. Immediately place the mold into a slurry of 2 parts

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crushed dry ice to 1 part 2-methylbutane until 80% of the OCT turns white. Place onto dry ice, and store at −80◦ C until sectioning. 5a. To section, allow each block to equilibrate for 5 min inside the cryostat (at approximately −20◦ C), adhere OCT block to the chuck with liquid OCT, cryosection into 4- to 30-μm coronal or sagittal (to visualize entire RMS) sections, and mount onto electrostatically charged Superfrost Plus glass slides. 6a. Allow the sections to dry for 30 min at room temperature. Store at −20◦ C or −80◦ C, for staining or RNA hybridization, respectively, up to at least 1 year.

To section using a vibratome We prefer sectioning on a vibratome when appropriate because consistent serial freefloating sections can be cut more easily and without the fluctuation in quality that the cryostat often displays. 3b. As an alternative to processing the tissue immediately, store brains in CMF-DPBS containing 0.1% sodium azide at 4◦ C to deter microbial growth and preserve tissue for up to a month before cutting with a vibratome. 4b. Place the brain into an appropriate mold and pour liquified low-melt agarose into the mold so that the entire brain is submerged. Allow the mold to harden at room temperature, then remove the agarose-embedded brain from the mold. Use of low-melting point agarose minimizes heat damage to the tissue.

5b. Mount samples onto a vibratome stand and section into 10- to 30-μm coronal or sagittal free-floating sections. 6b. Store sections in CMF-DPBS containing 0.1% sodium azide until ready to immunostain.

CHARACTERIZING hNPC Throughout expansion and long-term maintenance, hNPC are periodically assayed for in vitro expression of known “stemness” markers. Human NPC should be analyzed by standard immunoassays to demonstrate sustained undifferentiated characteristics and cellular morphology. Prior to transplantation, a subset of hNPC are set aside and assayed for the following markers (see below) in multiple combinations to confirm morphologically relevant expression profiles exist.

BASIC PROTOCOL 10

The fate of donor cells in the CNS can be assessed in vivo by dissecting the brains and processing by standard immunohistochemical methods for lineage-specific antigens such as nestin/vimentin for hNSCs; GFAP and s-100-Beta for astrocytes; and NeuN, NFM70/200, and synaptophysin for mature neurons. Antibodies to the human nuclear antigen (HuNuc) can be used to co-label hNSCs for donor confirmation. The functionality of donor cells in the brain can also be assessed by classic electrophysiological methods ex vivo.

Undifferentiated hNPC Healthy proliferative Ki-67+ (Scholzen and Gerdes, 2000; Ab available from Abcam, use at 1:200 dilution) or PCNA+ (Hall and Levison, 1990; Hall et al., 1990; Ab available from Santa Cruz Biotechnology, use at 1:100 dilution) undifferentiated hNPC express the nuclear transcription factor Sox-2 (D’Amour and Gage, 2003; Komitova and Erickson, 2004; Baer et al., 2007; Ab available from Santa Cruz Biotechnology, use at 1:100 dilution), musashi-1/2 (Chan et al., 2006; Ab available from Abcam, use at 1:100 dilution), the filamentous cytoplasmic proteins Nestin (Ab available from Chemicon, use at

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1:400 dilution; Fig. 2D.3.11A,B,E-J) and Vimentin (Ab available from Chemicon, use at 1:400 dilution), and the surface protein LexA/SSEA-1 (Ab available from Chemicon, use at 1:250 dilution) in cellularly appropriate locations (Temple, 2001; Pixley and de Vellis,1984; Pixley et al., 1984a,b; Hockfield and McKay, 1985; Lendahl et al., 1990; Dahlstrand et al., 1992; Zimmerman et al., 1994; Garcia et al., 2004). In addition, fetal hNPC also express glial fibrillary associated protein (GFAP; Ab available from DAKO, use at 1:400 dilution) in morphologically appropriate (nonprotoplasmic) locations, highly correlating with Nestin expression (Fig. 2D.3.11. E,G-J), consistent with rodent data (Doetsch et al., 1999; Laywell et al., 2000; Imura et al., 2003), suggesting that GFAP also labels human fetal NPC or that astrocytes may be hNPC at specific times during early development. It is critical that these markers be utilized in combination to confirm the stem/precursor phenotype of donor cells before transplantation. In addition to the classic stem cell markers, we have also confirmed the expression of radial glia (RG) associated brain lipid binding protein (BLBP; Feng et al., 1994; Feng and Heinz, 1995; Ab available from Chemicon, use at 1:350 dilution), which has been linked to fetal NSC in vivo (Fig. 2D.3.12; Garcia et al., 2004; Malatesta et al., 2000; Hartfuss et al., 2001; Alvarez-Buylla et al., 2001, 2002; Miyata et al., 2001; Noctor et al., 2001, 2002; Gotz et al., 2002; Gregg et al., 2002; Doetsch, 2003; Malatesta et al., 2003; Goldman, 2003; Gotz, 2003; Gregg and Weiss, 2003; Noctor et al., 2004, 2008; Merkle et al., 2004; Gotz and Bard, 2005; Merkle and Alvarez-Buylla, 2006; Merkle et al., 2007). In addition, a small population of bipolar migratory cells express the microtubuleassociated protein doublecortin (DCX; Fig. 2D.3.12E,F; Gleeson et al., 1999; Francis et al., 1999; Friocourt et al., 2003; Ab available from Santa Cruz, use at 1:400 dilution) and the membrane-bound polysialylated neural cell adhesion molecule (PSA-NCAM; Hu et al., 1996; Curtis et al., 2007; El Marouf and Rutishauser, 2008; Rutishauser, 2008; Burgess et al., 2008; Ab available from Chemicon, use at 1:250 dilution). Furthermore, some later-passage hNPC populations also express the intermediate filament protein, beta-3-tubulin (Tuj-1; Fig. 2D.3.11F; Caccamo et al., 1989; Geisert and Frankfurter, 1989; Lee et al., 1990; Menezes and Luskin, 1994; Menezes et al., 1995; Ab available from Covance Research, use at 1:400 dilution) and colocalizes with GFAP (Rakic, 1972; Sidman and Rakic, 1973; Levitt and Rakic, 1980; Levitt et al., 1981, 1983). Although beta-3-tubulin has been described as an early marker for immature neuron ally–restricted NPCs in rodents, these hNPC continue to self-renew and maintain multipotency in vitro and in vivo. Furthermore, they are phenotypically and behaviorally indistinguishable from early-passage predecessors. As a result, assuming the same stemness profile for human cells may not be entirely appropriate, and this should be taken into account when assessing hNPC fate. For example, expression of GFAP or beta-3-tubulin alone may not be entirely sufficient to assume loss of multipotency or regional differentiation into neurons and astrocytes, respectively. Instead, we favor a mode of characterization based on both morphology and multiple marker comparison to appropriately assess stem cell fate both before and after transplantation.

Long-Term MAN Growth and Characterization of NPCs

It is well known that cell lines accumulate in vitro artifacts in response to long-term artificial cell culture environments (Doetsch et al., 2002; Pollard et al., 2008). Whether the trends we see in vitro appropriately mimic the in vivo nature of hNPC remains to be determined. Certainly, it stands to reason that hNPC share many developmental markers in common with their rodent counterparts; however, it is not beyond the scope of reason to assert that perhaps some of the key players involved in human neural developmental processes may be differently regulated spatiotemporally from the corresponding processes in lower-order mammals. We have verified these findings by both standard immunocytochemistry and western blotting in three hNPC lines ranging from 10 to 22 weeks (from fertilization date), further suggesting that hNPC are not homogenous populations

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Figure 2D.3.11 hNSC express classic neural stemness genes. Human NSC (HFB-2050) express nestin (A,B) and Tuj-1 (C,D) uniformly when plated as aggregates on poly-D-lysine-coated tissue culture–treated flasks (blue = DAPI). Cells rapidly attach and begin to elongate, sending processes throughout the culture. (A) and (D) were processed 24 hr after plating, while (B) and (C) represent the more immature morphology seen after 12 hr of culture. DAPI+ nuclear morphology indicates active mitosis and sustained proliferation at both time points. In addition, nestin+ hNSC (green) also express GFAP (red) (E,H,J), typically in inverse proportion to each other. For example, a cell with high GFAP expression will also express nestin, but at a much lower level, whereas a highly nestin+ cell will express GFAP at a lower level. Interestingly, both cell types are intimately interwoven within each other, forming a meshwork of migratory cells (pictured here on polyL-ornithine/fibronectin-coated tissue culture–treated flasks). Furthermore, nestin+ hNSC also express Tuj-1 (F) throughout most of the cytoplasm. Typically, nestin expression is highest surrounding the nuclear box, whereas Tuj-1 expression is greatest at the feet of meandering processes. Expression patterns are typically opposite of each other, and tend to colocalize in the middle of the cell’s architecture, similar to nestin and GFAP coexpression. These filamentous proteins may play distinct roles at their specific positions within the stem cell, conferring or coordinating cell polarity within the in vitro microniche (G,I = phase contrast).

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Figure 2D.3.12 hNPC exist as heterogeneous populations resembling the SVZ niche. In addition to the classic NSC marker proteins, some cells also highly express brain lipid binding protein (BLBP) (A-H) or doublecortin (DCX) (E,F). Cells that highly express BLBP typically assume an astrocyte-like star morphology and have one or two long meandering processes with a highly arborized cell body, resembling radial glia. In contrast, most cells that do not express BLBP highly or have radial glia morphology express DCX and highly resemble the migratory transit amplifying cells found in vivo within the subventricular zone NSC niche. Images were taken from hNSC (HFB-2050) plated on Matrigel-coated tissue culture plates and cultured for 3 days in vitro to induce attachment and spreading. Of note, extracellular matrix components and growth factors found in Matrigel have a profound impact on cell morphology and may affect the differentiation profile of cultures over time. It is likely that these cultures have started an initial differentiation process and may not retain all stemness properties. Nonetheless, the heterogeneity of hNSC cultures is evident as 72 hr is probably not sufficient time to considerably differentiate hNSC.

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of identical stem cells, but rather highly dynamic heterogeneous populations (Kukekov et al., 1999; Laywell et al., 2002; Suslov et al., 2002; Steindler et al., 2003; Chen et al., 2006) of neural progenitors. Clonal analysis assays and in-depth time-lapse fluorescence video microscopy are currently being used to determine the specific lineage relationships involved. Whether these distinct phenotypic outliers represent bona fide stem cells or restricted progenitors that have been reprogrammed in response to mitogenic signaling cues (Kondo and Raff, 2000) remains to be determined. Certainly, the new wave of research dedicated to studying induced pluripotent stem cells (iPS) suggests that this process is much easier to induce than previously appreciated (Takahashi and Yamanaka, 2006; Takahashi et al., 2007 a,b; Maherali et al., 2007; Wernig et al., 2007; Okita et al., 2007; Meissner et al., 2007; Yu et al., 2007; Nakagawa et al., 2008; Park et al., 2008; Brambrink et al., 2008; Wernig et al., 2008; Stadtfeld et al., 2008; Shi et al., 2008; Kim et al., 2008; Maherali et al., 2008; Maherali and Hochedlinger, 2008a,b). Therefore, it is only through careful phenotypic characterization that we can begin to understand the nature of hNPC in vitro. Until better methods and markers are discovered to accurately access cellular identity, we are forced to apply the borrowed phrase, “It’s hard to define, but I know it when I see it” (Morrison et al., 1997).

Differentiated hNPC In addition to the classical stemness markers, cells may be differentiated into neurons, astrocytes, or oligodendrocytes by a variety of methods and assayed for lineage-specific differentiation markers. Specific methods for in vitro differentiation are detailed elsewhere (Wakeman et al., 2009; Johe et al., 1996; Hsieh and Gage, 2004; AndroutselisTheotokis et al., 2008). Differentiated cells are fixed in 4% cold PFA, stained with the appropriate antibodies by standard protocols, and analyzed by indirect immunofluorescence for the expression of pro-neuro/gliogenic markers. To determine whether hNPC are capable of giving rise to neurons (after allowing at least 3 weeks of in vitro maturation), the cells are stained for an extensive panel of pro-neuronal markers—first (immature): doublecortin (Santa Cruz, dilute 1:500), b-III-tubulin (Chemicon, dilute 1:200), Pax6 (Covance, dilute 1:400), Ptx3 (Abcam, dilute 1:500, or R&D Systems, dilute 1:400), Lmx1a/b (Santa Cruz, dilute 1:200), Gbx1/2 (Santa Cruz, dilute 1:250), Ngn1/2/3 (Santa Cruz, dilute 1:250), and then more mature neuronal phenotypes: PSA-NCAM (Chemicon, dilute 1:100), high-molecular-weight neurofilament (Boehringer, dilute 1:150), tau (Sigma, dilute 1:400), NeuN (Chemicon, dilute 1:50), MAP-2 (Sigma, dilute 1:200), synaptophysin (Sigma, dilute 1:500), calbindin (Sigma, dilute 1:400), calretinin (Chemicon, dilute 1:500), Mash1 (BD Biosciences, dilute 1:100), Msx1 (Abcam, dilute 1:500), En1 (Iowa Developmental Hybridoma Bank, dilute 1:50), Girk2 (Alomone Laboratories, dilute 1:500), Nurr-1 (Santa Cruz, dilute 1:500), TH (PelFreeze Biologicals, dilute 1:500, or primary rabbit antiserum, Eugene Tech, dilute 1:3000), DAT (rabbit, Affinity Bioreagents, dilute 1:500), VMAT2 (PelFreeze, dilute 1:500), AADC (Chemicon, dilute 1:1000; dopaminergic), GABA (Sigma, dilute 1:5000 or rabbit polyclonal antibody for GAD65/67, Chemicon, dilute 1:1000), ChAT (Chemicon, dilute 1:400; cholinergic), 5-HT (Sigma, dilute 1:50 or rabbit polyclonal, Calbiochem;, dilute 1:1200), as well as the radial glial, BLBP (Chemicon, dilute 1:1000), A2B5 (R&D Systems, dilute 1:400), the mature astroglial, GFAP (polyclonal rabbit, Dako-Patts, dilute 1:1000, Sternberger Monoclonals, dilute 1:200, or Chemicon, dilute 1:200), S100B (Abcam, dilute 1:500), glutamate transporters Glast/GluT-1/EAAT1 (Chemicon, dilute 1:400, or Santa Cruz, dilute 1:500) and GLT1/EAAT2 (Chemicon, dilute 1:400, or Santa Cruz, dilute 1:500), and oligodendrocyte fate, MBP (Chemicon, dilute 1:200), O4/O1 (Chemicon, dilute 1:200), CNPase (Chemicon, dilute 1:200), otx2 (Santa Cruz, dilute 1:100), and RIP (Chemicon, dilute 1:400).

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

5-Bromo-2 -deoxyuridine (BrdU), 1000× stock Prepare a 20 mM (1000×) stock solution by resuspending 60 mg 5-bromo-2 deoxyuridine (BrdU; Sigma, cat. no. 59-14-3) lyophilized powder in 10 ml DMSO or water, with heating (i.e., 60 mg BrdU into 10 ml sterile water). Vortex thoroughly until fully dissolved, filter sterilize through a 0.2-μm filter, aliquot 100 μl per sterile microcentrifuge tube, and store at –20◦ C for up to 6 months. As an alternative, utilize the BrdU Labeling and Detection Kit I, (Roche, cat. no. 1296736). The BrdU is supplied as a 10 mM (1000×) sterile solution in PBS. Note the concentration difference between brands.

hNPC freezing medium 20 ml NB-B-27 complete medium (see recipe; 40% v/v final) 25 ml fetal bovine serum (FBS; Invitrogen, cat. no. 16140-071) 5 ml dimethylsulfoxide (DMSO; Sigma, cat. no. D-2650) Filter sterilize using a 0.2-μm DMSO-Safe Acrodisc syringe filter (Pall, cat. no. 4433) Store up to 3 weeks at 4◦ C Hibernate-E medium, complete Prepare Hibernate-E medium (Brain-Bits; http://www.brainbitsllc.com/) supplemented with: 2% (v/v) B-27 supplement without vitamin A (Invitrogen, cat no. 12587-010) 8 μg/ml heparin 2 μl/ml Normocin After filter sterilization supplement with: 20 ng/ml basic fibroblast growth factor (bFGF; Millipore, cat. no. GF003) 10 ng/ml leukemia inhibitory factor (LIF; Millipore, cat. no. GF003) Store up to 3 weeks at 4◦ C Hibernate-E is a basal medium that supports short-term hNPC maintenance at ambient carbon dioxide levels (U.S. Patent 6,180,404).

NB-B-27 complete growth medium For 50 ml: 48.4 ml Neurobasal medium (Invitrogen, cat. no. 21103-049; 97% v/v final) 1 ml B-27 supplement without vitamin A (Invitrogen, cat no. 12587-010; 2% v/v final) 0.5 ml GlutaMAX (Invitrogen, cat. no. 35050-061; 1% v/v final) 400 μg heparin (Sigma, cat. no. H-3149; 8 μg/ml final) 2 μl/ml Normocin (InvivoGEN) or other concentration as specified in protocol (optional) Filter sterilize, then add growth factors (bFGF, EGF, LIF) as specified in protocol Paraformaldehyde, 4% (w/v)

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Under a chemical fume safety hood, add 40 g of paraformaldehyde to 860 ml of distilled water, heat to 55◦ C, and stir until uniformly dispersed. Do not exceed 55◦ C, as paraformaldehyde is highly unstable at higher temperatures and will generate toxic by-products. Add several drops of 1 M NaOH until the solution is fully dissolved and cleared (clearing will not occur without adding NaOH). continued Current Protocols in Stem Cell Biology

When solution clears, add 100 ml of 10× DPBS (Invitrogen, cat. no. 14200-067) and adjust total volume to 1 liter. Adjust pH to 7.6 with NaOH or HCl and cool to 4◦ C. Prepare this solution fresh, and modify volumes to make appropriate amounts. Alternatively freeze in aliquots at −20◦ C for later use. Do not freeze/thaw more than once. We strongly suggest preparing PFA fresh at the time of use for optimal results.

COMMENTARY Background Information The great neuroanatomist Ram´on y Cajal wrote, “In the adult centers the nerve paths are something fixed, ended and immutable. Everything must die, nothing may be regenerated” (Ram´on y Cajal, 1928; Ram´on y Cajal and May, 1959). This observation, based on the primitive methods of the time, held up as neurodevelopmental dogma for centuries. It wasn’t until Joseph Altman and Michael Kaplan’s classic autoradiographic experiments using tritiated thymidine (Altman, 1962a,b, 1963; Altman and Chorover, 1963; Altman and Das, 1965a,b, 1966) that neurobiologists even considered rethinking the notion of adult neurogenesis (Allen, 1912; Messier et al., 1958; Messier and Leblond, 1960; Smart, 1961; Smart and LeBlond, 1961; Kaplan and Hinds, 1977), let alone embraced it as an intrinsic process active throughout adulthood until death. In the past quarter century, it has become undeniably clear that neural stem/progenitor cells reside within the developing embryonic, neonatal, and adult songbird (Goldman and Nottebohm, 1983), rodent (Alvarez-Buylla et al., 2002; Snyder et al., 1992; Morshead et al., 1994; Weiss et al., 1996; Johanson et al., 1999), monkey (Gould et al., 1998, 1999a,b; Kornack and Rakic, 2001a,b), and human forebrain (Merkle et al., 2007; Curtis et al., 2007; Sanai et al., 2004; Howard et al., 2006; Quinones-Hinojosa et al., 2006, 2007; Sanai et al., 2007), primarily lining the posterior to anterior subventricular zone (SVZ; Doetsch et al., 1999; Merkle et al., 2004; Menezes et al., 1995; Lois and Alvarez-Buylla, 1993; Luskin, 1993; Lois and Alvarez-Buylla, 1994; Rousselot et al., 1995; Luskin et al., 1997) of the lateral ventricular walls and within the subgranular zone of the hippocampal dentate gyrus (Ray and Gage, 2006; Ray et al., 1993; Gage et al., 1995a). These cells can be derived from various regions of the brain, with limited capacity, where they exist naturally as relatively quiescent populations of stem and progenitor cells (Palmer et al., 1995) within the complex microenvironment of a highly dynamic, tightly junctioned, neurogenic niche

(Sanai et al., 2007; Alvarez-Buylla and Lim, 2004; Verdugo and Alvarez-Buylla, 2006; Lim et al., 2007). During mammalian CNS development, hNSC undergo an initial expansion phase of symmetric divisions followed by nonsymmetric divisions and extensive migration in accordance with electrical stimulation (Deisseroth et al., 2004; Spitzer, 2006) and chemical (Ghashghaei et al., 2007) guidance cues. Differentiation is characterized by stages of neurogenesis followed by gliogenesis (Qian et al., 1997, 1998, 2000; Namihira et al., 2009), differentiating in temporal waves of first neurons, then astrocytes and oligodendrocytes, to shape and form the mature human brain (Levison and Goldman, 1993; Levison et al., 1993; Menn et al., 2006). In the adult, NPC continue to proliferate and migrate by a combination of tangential and chain migration from the SVZ through the rostral migratory stream (RMS) into the olfactory bulbs (OB; Lois et al., 1996; Doetsch and Alvarez-Buylla, 1996), generating new neurons and glia (Kuhn et al., 1996; Goldman et al., 1997; Petreanu and AlvarezBuylla, 2002; Carleton et al., 2003; Lledo et al., 2006) as an active pool to replace or restore homeostasis to the aging or injured brain (de Chevigny et al., 2008; Lim et al., 2007; Nait-Oumesmar et al., 1999; PicardRiera et al., 2002; Lie et al., 2004; Parent et al., 2006; Leung et al., 2007; Hellstrom et al., 2008). In addition, cultured NSC/NPC can be directed in vitro and in vivo to give rise to all three neuroectodermal lineages (Arsenijevic et al., 2001a; Johe et al., 1996; Gage et al., 1995b; Kirschenbaum et al., 1994; McKay, 1997; Levison and Goldman, 1997; Murray and Dubois-Dalcq, 1997; Pincus et al., 1998; Takahashi et al., 1999; Rao, 1999; Brannen and Sugaya, 2000; Dietrich et al., 2002; Riaz et al., 2004; Scheffler et al., 2005; Christophersen et al., 2006; Pistollato et al., 2007; Rao et al., 2008), as well as a variety of intermediate cellular phenotypes (Markakis et al., 2004) when presented with the appropriate signaling cues. These cells may act by a variety of mechanisms, either by providing

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potential new raw material for regenerating the damaged CNS or by rescuing the endogenous nervous system through secondary neuroprotection mechanisms (Pluchino et al., 2005a), thereby modulating the host microenvironment (Madhavan et al., 2005, 2006, 2008), conferring a return to baseline nonpathological stasis (Bjugstad et al., 2005). In this manner, neurological function may be restored through transplantation therapies by either directly integrating and replacing host neural circuitry, or, secondarily, by rescuing and restoring the endogenous host milieu through growth factor or neurotransmitter paracrine signaling (Brustle and McKay, 1996; Park et al., 2002a; Svendsen and Langston, 2004; Tai and Svendsen, 2004; Emsley et al., 2005; Pluchino et al., 2005a,b; Schwartz, 2006). Moreover, multipotent hNSC have been shown to display low immunogenicity (Mason et al., 1986; McLaren et al., 2001; Odeberg et al., 2005), readily express foreign transgenes (Flax et al., 1998; Ostenfeld et al., 2002a; Wu, P. et al., 2002; Park et al., 2003; Behrstock and Svendsen, 2004; Kim, 2004; Klein et al., 2005; Behrstock et al., 2006; Capowski et al., 2007; Roy et al., 2007; Suzuki et al., 2007; Ebert et al., 2008), and inherently home to sites of pathological insult (Aboody et al., 2000; Park et al., 2002b; Ourednik et al., 2002; Imitola et al., 2004; Park et al., 2006). Traditionally, human neural stem cells (hNSC) have been operationally defined (Weissman et al., 2001; Anderson, 2001; Seaberg and van der Kooy, 2003; Parker et al., 2005; Navarro-Galve and Martinez-Serrano, 2006) by two cardinal criteria, first, the ability to self-renew indefinitely by division into two identical (symmetric) or nonidentical (asymmetric) daughter cells, and, second, to maintain the multi-potential capacity to differentiate into all three neuroectodermal CNS lineages: neurons, astroglia, and oligodendrocytes in vivo (Gage et al., 1995a; Brustle et al., 1997; Lundberg et al., 1997; Winkler et al., 1998; Zigova et al., 1998; van der Kooy and Weiss, 2000; Weissman et al., 2001; Buchet et al., 2002). Asymmetric daughter cells that are not identical to the parental hNSC have a reduced stemness and are therefore referred to as intermediate or transitamplifying neural progenitor cells (Doetsch et al., 1999; Merkle and Alvarez-Buylla, 2006; Verdugo and Alvarez-Buylla, 2006; Doetsch et al., 1997; Garcia-Verdugo et al., 1998; Lim et al., 2008; for reviews see Kemperman, 2006; Slack, 2008). In vertebrates, NSC self-renewal

can be viewed as a property of the entire population as opposed to just the single cellular entity, in that self-renewal is thought of more globally as the capacity to maintain the absolute number of neural stem cells in a given area at a steady level throughout time (Gritti et al., 2003). This more relaxed definition allows for fluctuations in absolute NSC population size, evenly balancing symmetric stem cell divisions with asymmetric differentiated progenitors as physiological conditions dictate. Although there may be no true NSC capable of self-renewing indefinitely throughout adulthood in vivo, NPCs seem to maintain prolonged self-renewal (Ravin et al., 2008) and regain their multipotentiality when exposed to mitogenic growth factors in vitro (Doetsch et al., 2002; Gabay et al., 2003). In addition, transplantation of SVZ-derived NPC suggests that they retain both migratory and differentiation capabilities when homotopically reintroduced into appropriate anatomical locations, but may only differentiate and fail to migrate when heterotopically positioned (Betarbet et al., 1996; Zigova et al., 1996; Hererra et al., 1999; Yang et al., 2000; Ourednik et al., 2001; Tamaki et al., 2002; Seidenfaden et al., 2006). The results demonstrate that there may be an intrinsic spatiotemporal program that determines the developmental potential of NPC. This apparent positional restriction suggests heterogeneity of hNPC as restricted populations of bipotent or unipotent glial and neural precursor cells in vivo; however, these finding do not exclude the possibility that precursor cells may reconfer multipotentiality in vivo (Ostenfeld et al., 2002b; Hitoshi et al., 2002; Parmar et al., 2002, 2003; Lepore et al., 2004; Kim et al., 2006; Kallur et al., 2006) upon long-term exposure to the local cellular milieu or complementary signaling of mitogenic growth factors from secondary expansion in vitro. Recent evidence from Cre-lox and retroviral lineage mapping studies has challenged the past notion that NPC become fate restricted before adulthood (Garcia et al., 2004; Doetsch et al., 1999; Malatesta et al., 2000; Hartfuss et al., 2001; Miyata et al., 2001; Noctor et al., 2001; Gotz et al., 2002; Doetsch, 2003; Malatesta et al., 2003; Goldman, 2003; Gotz, 2003; Noctor et al., 2004; Merkle et al., 2004; Gotz and Barde, 2005; Noctor et al., 2008; Weissman et al., 2003; MartinezCerdeno et al., 2006). These data have led to three major changes in the tenets of developmental neurobiology (Merkle and

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Alvarez-Buylla, 2006; Malatesta et al., 2008). First, adult NSC are of glial origin, but are not fate restricted and can therefore give rise to all three central nervous system (CNS) cell subtypes. Second, embryonic, fetal, and adult NSC are uniformly lineage related through radial glia (RG), and, third, fetal NPC actually divide asymmetrically to increase the number of progeny they generate via symmetrically dividing intermediate NPCs (Alvarez-Buylla et al., 2001; Tramontin et al., 2003; Rakic, 2007). The revised unified model demonstrates the lineal transition from neuroepithelium to radial glia and eventually adult astrocyte-like NPC and may help explain the relative heterogeneity that NPC display throughout embryonic and fetal development into adulthood. Furthermore, the unified theory proposes that functional CNS stem cells will display heterogeneous phenotypes throughout neural development depending on temporal and spatial cues, suggesting that the nature of in vivo NSC/NPC is highly dynamic. Moreover, the relative heterogeneity and plasticity of the in vivo neurogenic niche suggests a similar component may exist within artificial in vitro dissociated stem cell culture preparations.

Critical Parameters and Troubleshooting To successfully expand and maintain hNPC, it is imperative to adhere to the following recommendations throughout all procedures to ensure maximum recovery. Mechanical shear forces can easily damage the extremely fragile cell membrane; therefore hNPC are always triturated gently and centrifuged at low speed for short periods of time to reduce cell death, increase the viability, and maximize the overall recovery of hNPC. Careful consideration should be made when removing any supernatant following centrifugation, so that the cell pellet is not lost. To remove supernatant, simply tilt the conical tube to a 45◦ angle and remove all but the lower 50 to 100 μl of supernatant directly above the cell pellet (∼1 mm below meniscus of supernatant). Furthermore, the MAN culturing assay requires precise coordination of timing and density to properly establish the webbed structure that supports extensive growth. We prefer uncoated tissue culture–treated 0.22-μm vented cap flasks to the standard petri-dish style culture vessels for expansion of undifferentiated hNPCs, due to the lower risk of contamination. It is also much easier to evenly scatter the cells in rectangular flasks Current Protocols in Stem Cell Biology

in comparison to circular well–style dishes and plates, which often result in the aggregation of large cellular clusters at the center of the well. Once cell-cell contact has been made, the separate aggregates prematurely merge into large irregular masses (Mori et al., 2006; Reynolds and Rietze, 2005; Singec et al., 2006; Jessberger et al., 2007; Ren et al., 2007; Marshall et al., 2007; Singec and QuinonesHinojosa, 2008) and adhere together through surface integrins and self-secreted extracellular matrix proteins (Flanagan et al., 2006; Jacques et al., 1998; Hynes, 2002; Campos et al., 2004; Leone et al., 2005; Mueller et al., 2006). Uneven distribution of single cells can quickly deter the formation of small NPC clusters, resulting in aggregation of hNPC into large globular masses. As a result, cells located at the periphery of the cluster will begin to differentiate (Svendsen et al., 1998). Furthermore, cells located at the central core may become necrotic (Svendsen et al., 1997a,b) from inadequate gas exchange and lack of nutrients required for normal metabolic activity (indicated by dark pigmentation within cellular masses under phasecontrast microscopy). Populations of hNPC with necrotic cells should be thoroughly dissociated to remove the cellular debris and rescue the proliferative precursors that remain. The overall result of the untimely premature dissociation is the loss of many hNPC, as most of the smaller cell clusters are extremely fragile and may be damaged by the extra round of enzymatic dissociation. Passaging of the MAN The dissociation process is the fundamental technique for successful hNPC culture and lies at the basis of every major procedure employed, from basic expansion and maintenance, to single cell cloning, genetic manipulation, labeling, and in vivo transplantation. Whether employing enzymatic techniques or modified salt solutions, specific emphasis is placed on optimizing recovery of viable progenitors throughout this fundamental procedure. Dissociation (passaging) of MAN hNPC cultures presents a unique dilemma due to the highly dynamic microtubule reorganization and migratory properties that hNPC exhibit. In order to preserve the integrity of these polarized progenitors, gentle release and reorganization of meandering growthcone extensions must be achieved to effectively dissociate the highly intertwined processes without damaging the integrity of the cell membrane. Achieving homogenous,

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high-viability, single-cell dissociation requires precise coordination of low-speed controlled trituration coupled with minimizing the amount of time progenitors spend incubating in enzymatic agents, so as not to destroy the fragile hNPC. Determining the precise ratio of these elements that best complements your specific hNPC line depends on a variety of factors including overall three-dimensional cell density and degree of adherence to the culture vessel. When MAN hNPC cultures become overgrown (>75% confluent), become inadvertently detached by mechanical stress following dissociation, or display the initial signs of necrosis (yellow to brownish colony cores), Basic Protocol 1 is employed to dissociate cells. Generally, we use the enzymatic agent Accutase to dissociate hNPC into single cells, as it is much more gentle with fragile hNPC than trypsin-EDTA and does not require chemical inactivation. Accutase is simply diluted 1:5 to 1:10 in basal growth medium followed by centrifugation to eliminate residual Accutase and remove cellular debris. Other dissociation agents such as Accumax, TrypLE, and collagenase may also be implemented provided that incubation times and inactivation steps are adapted as necessary. The remainder of the procedure would remain the same. In addition, the nonenzymatic, Hanks’-based cell dissociation buffer (CDB) can also be employed for applications such as FACS analysis where extension processes and surface receptors must remain intact. CDB slowly detaches adherent hNPC, so the delicate neurite processes tend to retract from each other more gently in comparison to Accutase, which can cleave delicate extension processes when used excessively. We do not generally utilize CDB for standard expansion due to the relatively long incubation times required to break hNPC clusters into single cells. In our hands, the partial lifting of adherent clusters with CDB often results in mass clumping and cell death, evidenced by large quantities of sticky DNA precipitates in solution. This process can require multiple rounds of treatment to separate and thoroughly detach and dissociate single cells from the remaining highly branched hNPC colonies. The additional mechanical trituration, centrifugation, and subsequent cell death leads to decreased recovery of viable cells compared to the quicker-acting Accutase. When using CDB, the procedure remains as follows but incubation times are increased up to 20 to 35 min to achieve a similar level of dissociation as Accutase.

In addition, chopping large spheres into smaller cellular clusters (Svendsen et al., 1998; Anderson et al., 2007) has been shown to be highly beneficial in comparison to single-cell dissociation so as not to destroy integral cellcell contacts critical for continued proliferation. We can obtain the same effect, creating the same small-size cellular clusters by decreasing the Accutase incubation time and inactivating the suspension before the cellular clusters are fully dissociated into single cells. A similar effect can be obtained though a slightly different approach, where single cells are replated at a higher density than typically employed to rapidly induce cell-cell contacts and early aggregation of small hNPC clusters through premature merging. Within 6 to 24 hr, small- to medium-size aggregates will form, similar in size to spheres created by mechanical chopping. Furthermore, single cells can easily be counted with a hemacytometer and accurately replated at known densities for accurate record keeping of sustained stem cell growth dynamics within cultures. Although it has been published that passaging cells with enzymes results in “high risk of high rates of cell death, lack of adherence, or differentiation” (Nethercott et al., 2007) as well as induction of karyotypic abnormalities, utilizing the procedures described here, we have been able to maintain behaviorally normal, karyotypically stable, undifferentiated forebrain hNPC (Villa et al., 2004; Foroni et al., 2007) as highly proliferative, multilayer adherent networks for >100 passages without marked senescence or phenotypic adaptation by means of enzymatic (Accutase) single-cell dissociation. It is our opinion that overall expansion rates and possibly time to senescence (Carpenter et al., 1999; Goyns and Lavery, 2000; Wright et al., 2006) can be greatly increased by simply improving the overall condition of hNPC during and after dissociation, regardless of the technique employed. The repetitive combination of mechanical shear stress from trituration, centrifugation, and osmotic shock simply provides more opportunities to destroy the fragile neural progenitors and ultimately results in a gradual decline in hNPC numbers. Furthermore, we speculate that as the gross number of actively mitotic progenitors decreases, the subsequent loss of paracrine signaling (Taupin et al., 2000; Toda et al., 2003; Agasse et al., 2004, 2006) between hNPC eventually falls below a threshold concentration, whereby the delimited hNPC culture no longer maintains the capacity to properly condition its own basal

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substrate and subsequently becomes quiescently static, undergoing an irreversible halt in paracrine/autocrine regulatory signaling. The ultimate result of such events is a small population of nonproliferative hNPC in severe crisis; these cells are not suitable for study and should be distinguished from their proliferative counterparts and discarded. We propose a model, whereby hNPC endterm senescence and proliferative potential is influenced by population density through “conditioned signaling” and can be controlled by manipulating various combinations of these factors. Moreover, in vitro human manipulation can play a huge impact on the overall health and success of cultures, impacting the combined intrinsic signaling cascades that govern the phenotype of hNPC. On a global scale, the ultimate capacity for longterm self-renewal and ability to generate extremely large quantities of undifferentiated neural precursors (Svendsen and Smith, 1999) may be vastly improved with minimal adaptation to currently employed procedures. We therefore posit that the potential for somatic hNPC therapy and diagnostics would best benefit by a paradigm shift in culturing techniques from low- to high-density adherent populations, paying special attention to the importance of re-establishing essential cell-cell contacts. Investigating these properties may restructure the current theory of in vitro populations of somatic hNPC as limited-capacity progenitors (Hayflick, 1968; Temple and Raff, 1986; Durand et al., 1998; Svendsen et al., 1998; Quinn et al., 1999; Palmer et al., 2001; van Heyningen et al., 2001) incapable of amassing the relatively large quantities of cells (like their embryonic counterparts) necessary for regenerative therapies (Gottlieb, 2002).

Anticipated Results The long-term expansion and continued maintenance of hNPC is a complex, highly dynamic process with many underappreciated intricacies. The procedures we describe here are intended as a general outline by which to adapt to the specific intricacies of your intended assay. Protocols can be adjusted according to the dynamics and behavior of each specific hNPC line, as individual cultures often vary highly in their specific dynamics and must be manipulated accordingly. Procedures may appear fairly clear-cut, but hNPC cultures are often highly variable in their composition and often deviate from the predictable nature of standard tissue culture. To accommodate for these changes, it may be necessary to alter standard

protocols on an impromptu basis to ensure the long-term stability of healthy hNPC cultures.

Time Considerations In general, every effort should be made to minimize time spent outside of normal proliferative conditions. It is extremely important to adhere to strict timing outlined in the procedures, particularly when establishing multilayer adherent network cultures. The initial adherence and expansion relies on controlled timing (3 to 4 days) for the even distribution of webbed hNPC. Alterations in duration of the procedure may result in unwarranted differentiation or apoptosis. In addition, the dissociation process should be optimized so that cells are not in contact with enzymes for long periods of time. Furthermore, many medium components are only stable for short periods of time; therefore, supplementation of basal medium is recommended every 48 to 72 hr to properly balance the formulation. All experiments involving human tissue must be approved by the appropriate institutional and/or national review boards and human tissue must be obtained with informed consent.

Acknowledgements D.R. Wakeman would like to thank Steven A. Wakeman and Pamela S. Burnett for constructive comments and support, as well as Ilyas Singec, Scott R. McKercher, Michael Marconi, Jean-Pyo Lee, and Kook I. Park for technical advice and procedural training. Funding for D.R.W. comes from (NIH/NIGMS T32 GM008666) UCSD Institutional Training Fellowship in Basic and Clinical Genetics, HHMI Med-Into-Grad Training Fellowship, American Society for Neural Therapy and Repair, and the American Parkinson’s Disease Association. Additional support was provided by the Stem Cell Center at the Burnham Institute for Medical Research (NIH P20 GM075059-03). The authors declare no conflicting or competing financial interest.

Literature Cited Aboody, K.S., Brown, A., Rainov, N.G., Bower, K.A., Liu, S., Yang, W., Small, J.E., Herrlinger, U., Ourednik, V., Black, P.M., Breakefield, X.O., and Snyder, E.Y. 2000. Neural stem cells display extensive tropism for pathology in adult brain: Evidence from intracranial gliomas. Proc. Natl. Acad. Sci. U.S.A. 97:12846-12851. Agasse, F., Roger, M., and Coronas, V. 2004. Neurogenic and intact or apoptotic non-neurogenic areas of adult brain release diffusible molecules that differentially modulate the development of

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subventricular zone cell cultures. Eur. J. Neurosci. 19:1459-1468.

McKay, R.D. 2008. Generating neurons from stem cells. Methods Mol. Biol. 438:31-38.

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Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S.,

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Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

UNIT 2D.4

Araceli Espinosa-Jeffrey,1 Dustin R. Wakeman,2, 3 Seung U. Kim,4 Evan Y. Snyder,3 and Jean de Vellis1 1

David Geffen School of Medicine at UCLA, Los Angeles, California University of California at San Diego, La Jolla, California 3 The Burnham Institute for Medical Research, La Jolla, California 4 University of British Columbia Hospital, Vancouver, British Columbia, Canada 2

ABSTRACT Here we document protocols for the production, isolation, and maintenance of the oligodendrocyte phenotype from rodent and human neural stem cells. Our unique method relies on a series of chemically defined media, specifically designed and carefully characterized for each developmental stage of oligodendrocytes as they advance from oligodendrocyte progenitors to mature, myelinating oligodendrocytes. Curr. Protoc. Stem Cell C 2009 by John Wiley & Sons, Inc. Biol. 10:2D.4.1-2D.4.26.  Keywords: neural stem cells r NSC r oligodendrocyte specification r oligodendrocyte maturation r lineage progression r oligospheres r neurospheres

INTRODUCTION In this unit, protocols are provided for the derivation, expansion, and maintenance of the oligodendrocyte (OL) phenotype from both rodent and human neural stem cells (NSC). This unique method utilizes chemically defined media, each formulated and carefully characterized for specific developmentals stages of OL as they advance from OL progenitors (OLP) to mature myelinating OL (Fig. 2D.4.1; Neman and de Vellis, 2008). By providing hNSC with the nutrients specifically required at a particular moment in OL development, our system allows for the propagation of OL at a desired stage from OLP to mature premyelinating OL. Therefore, lineage progression can be manipulated by controlling the duration of a given developmental stage as needed, in a more “natural” manner, and without using gene transfer (Park et al., 2002b; Kim, 2004; M¨uller et al., 2006; Ahn et al., 2008), cocultures, or undefined substrates such as cell line–derived conditioned medium or animal serum.

Preparation of embryonic neural stem cells (NSC) The methodology described in this unit can be used to isolate and derive NSC lines from various species. Specific methods for the derivation of human NSC are detailed elsewhere (Svendsen et al., 1999; Villa et al., 2000; Palmer et al., 2001; Schwartz et al., 2003; Kim et al., 2006; De Filippis et al., 2007; Kim et al., 2008; Wakeman et al., 2009; also see UNITS 2D.2 & 2D.3). ISOLATION OF RODENT NEURAL STEM CELLS In this protocol, 1 to 14 embryos can produce a successful preparation because stem cells can be propagated many times to obtain the desired yield.

BASIC PROTOCOL 1

Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.4.1-2D.4.26 Published online September 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d04s10 C 2009 John Wiley & Sons, Inc. Copyright 

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stem cells embryonic

neural

NKX2.2 PSA-NCAM Nestin, VIM CD133, Pax6

OL progenitor

PDGFR Olig 1 Olig 2 NG2, GD3 A2B5, Sox9 NKX6, 1 NKX2.2

pre-OL

O4 Tf CNP GPDH PDGFR Olig 1 Olig 2

immature OL

mature OL

RIP GC O1 O4 Tf CNP GPDH

MBP, PLP GSTII, MOG RIP GC O1 O4 Tf CNP GPDH

GDM

OLDEM

SSeA-1 Oct4 Sox9 STM

OSM 20 hr

OSM 1,2 day

Figure 2D.4.1 Oligodendrocyte specification and lineage progression. Oligodendrocytes undergo sequential morphological changes as they develop from uncommitted NSC to a committed OLP and acquire characteristics inherent in a functional OL. The list of OL markers below each developmental stage is not exhaustive but represents frequently used markers to identify OLs and their developmental stage. Media (also see Reagents and Solutions): stem cell medium (STM; Espinosa-Jeffrey et al., 2002); OL specification medium (OSM; Espinosa-Jeffrey et al., 2002); glia defined medium (GDM; Espinosa de los Monteros and de Vellis, 1996); OLDEM (Espinosa de los Monteros et al, 1988, 1997). Modified from Arenander and de Vellis (1995).

Materials One timed-pregnant, embryonic day 14 to 16 (ED14 to ED16) Sprague-Dawley rat (Charles River Laboratories) Basal stem cell medium (STM-II; see recipe) supplemented with 1% (w/v) BSA (Sigma, cat no. A-3156) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) Complete stem cell medium (STMIIc; see recipe)

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Dissection instruments, sterile (refer to numbers in Figure 2D.4.2): No. 1. Mayo scissors (Fine Science Tools, cat. no. 14010-17) No. 2. Lister scissors (Fine Science Tools, cat. no. 14131-14) No. 3. blunt-pointed forceps (Fisher, cat. no. 08-887) No. 4. iris scissors (Fine Science Tools, cat. no. 14060-09) No. 5. Moria iris forceps (Fine Science Tools, cat. no. 11373-12) No. 6. Dumont #7 forceps (Fine Science Tools, cat. no. 11297-10) No. 7. 140-μm and 230-μm sieves (Cellector, E-C Apparatus Corp.; http://www.thermo.com)

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No. 8. 20-ml syringe (Kendall, cat. no. 520673; http://www.kendallhq.com) No. 9. 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 100 × 15–mm petri dish (bacterial grade, non TC-treated; BD Falcon, cat. no. 351029) 50-ml and 15-ml conical tubes Centrifuge (e.g., IEC Clinical) 100-mm anti-PSA-NCAM coated dishes (Support Protocol 1) 37◦ C, 4.5% CO2 incubator (adjustable to 5% if growth is slow), 95% humidity Additional reagents and equipment for isoflurane anesthesia of the mouse (UNIT 2A.5), assessing cell viability (Support Protocol 2), and counting cells using a hemacytometer (UNIT 1C.3) NOTE: All dissection instruments, plasticware, and glassware must be sterile. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals.

Collect the embryo brains 1. Prepare the work area and sterile tools in a biosafety hood (Fig. 2D.4.2). 2. Euthanize the rat by isoflurane inhalation (UNIT 2A.5). 3. Dissect and remove the uterus. Collect the placenta-containing embryos and place in basal stem cell medium containing 1% BSA at room temperature in a non-tissueculture-treated 100-mm-diameter petri dish. 4. Remove the embryos from their placenta. Place them in a 100-mm petri dish containing PBS at room temperature. Remove the cerebellum from each embryo.

7 8 filter 9 PBS 3

5

needle

Hank’s 6

4

2

1

Figure 2D.4.2 Instruments required for dissection. (1) Scissors for decapitation; (2) scissors to cut the head skin to expose the skull; (3) forceps to hold the head in place as you cut the skin and cut the skull cartilage with scissors (4) to expose the brain. Some users prefer the curved scissors (2) instead of (4). Use the same scissors to transfer the brain to the petri dish containing PBS. Forceps (5) and (6) are to hold the brain in place and remove the meninges, respectively. After removal of the meninges, place the brains in HBSS while dissecting the rest of the brains. A filter mesh (7) is used to filter the cell suspension after dissociation. A 20-ml sterile syringe (8) and sterile 18-G dissociation needle (9) are used to dissociate the cells.

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5. Dissect out the brain of each embryo and place in STM-II complete (STMIIc) medium at room temperature in a 100-mm non-tissue-culture-treated petri dish. 6. Separate the cortex from the rest of the brain and remove the meninges with forceps. Once devoid of meninges, combine cortex and pons (i.e., the rest of the brain).

Isolate the cells 7. Combine the tissue of all the brains without meninges in a 15-ml conical tube. Mechanically dissociate with the 18-G needle (attached to a 20-ml syringe) by gently aspirating the brain pieces (10 times) and releasing the suspension slowly with the needle against to the wall of the tube (try to minimize foaming). 8. Centrifuge 8 min at 450 × g, room temperature. Recover the supernatant with the cells in suspension and transfer to a 15-ml tube. 9. Add 2 to 4 ml of STMIIc to the chunks left over in the dissociation tube and dissociate again five to eight times. In place of steps 7 to 9, you can dissociate the cells for 2.5 min in a Stomacher 80 (Seward; http://www. brinkmann.com).

10. Filter the suspension of dissociated cells first through the 230-μm sieve and then through the 140-μm sieve to remove cell clusters. 11. Rinse the two sieves sequentially with 2 ml STM-II medium containing 1% BSA at room temperature, and add this medium to the tube containing the cells. 12. Collect the cells by centrifugation 8 min at 450 × g, room temperature. 13. Gently discard the supernatant. 14. Resuspend the cell pellet in 4 ml of complete stem cell medium (freshly prepared), and gently dissociate the pellet with the syringe and needle by aspirating it up and down twice.

Initiate the cultures 15. Assess cell viability (Support Protocol 1), count cells using hemacytometer (UNIT 1C.3), and plate onto fresh PSA/NCAM–coated dishes (2 × 106 cells/100-mm dish in 7 ml of STMIIc medium). 16. Incubate plated cells overnight at 37◦ C with 4.5% CO2 and 95.5% humidity. Younger cells do not yet express PSA-NCAM and will remain floating as small clusters, whereas the older cells will attach overnight.

Collect conditioned medium (CM) 17. On the next day, transfer the nonattached cells to a 15-ml conical tube, pellet cells by centrifugation for 8 min at 500 × g, room temperature, and remove and save the conditioned medium (CM). ◦

18. Collect, filter, and save the conditioned medium at 4 C for immediate use (or frozen for later use). CM is an excellent supplement to start NSC cultures from frozen stocks. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

19. Allow attached cells (see step 16) to grow to 70% to 90% confluency. 20. Resuspend the pellet from step 17 and dissociate in 4 ml of fresh STMIIc medium by passing through a 14-G needle eight times. Bring the volume to 8 ml with conditioned medium and plate on additional anti-PSA-NCAM coated plates. Alternatively, to dissociate the cells, place 1.0 ml of the cell suspension in a sterile 75-ml Erlenmeyer flask in 25 ml of STMIIc, and incubate with shaking at 37◦ C (Fig.2D.4.3).

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E-16 cell suspension (mechanical dissociation) STMIIc

plate on anti-PSA-NCAM (coated dishes)

supernatant can be dissociated and re-panned

overnight

recover attached cells replate on fresh anti-PSA-NCAM

overnight

2-D cultures

NSC studies 3-D cultures

2-D

3-D attached on poly-D-lysine (well plates, chambers, dishes)

propagation

NSC

neurospheres

cryopreseration

neurospheres

Figure 2D.4.3 Rodent neural stem cell preparation. Following dissection, the cell suspension is plated on antiPSA-NCAM antibody–coated dishes and allowed to adhere. The process can be performed repeatedly to increase the numbers of neural stem cells, as two-dimensional cultures or three-dimensional “sphere” cultures (shown on the left side of the diagram). Every time cells are propagated, use anti-PSA-NCAM-coated dishes. Alternatively, cells can be propagated and immediately used for cell culture experiments (as shown on the right side of the diagram). While we prefer to use committed OL progenitors for cell transplants, other investigators also use uncommitted progenitors for grafting. Somatic Stem Cells

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21. Feed all cells every other day by removing 1/3 of the culture medium and adding the same volume of fresh STMIIc. 22. Switch the cells from 4.5% to 5.0% CO2 only if the cells are growing slowly. Leave them at 4.5% if the color of the medium stays red/orange. 23. Optional: To assess the phenotype of these cells, perform immunocytochemistry (Espinosa et al., 2002). SUPPORT PROTOCOL 1

PREPARATION OF ANTI-PSA-NCAM-COATED DISHES FOR SELECTING NSC BY IMMUNOPANNING The following method was developed based on published work (Wysocki and Sato, 1978; Williams and Gard, 1997) to isolate the rodent NSC population from the other cell populations in the brain during initial plating. We also use anti-PSA-NCAM coated dishes to propagate rodent NSC in two-dimensional cultures (Espinosa-Jeffrey et al., 2002). Please refer to the literature for specific methods on the selection of human NSC during primary derivation (Wakeman et al., 2009). We have chosen immunopanning as opposed to flow cytometric cell sorting because we find that cell survival approaches 100% when selecting the desired cell type via immunopanning. We know from both the experience of other scientists and our own that the survival rates are never this high when using FACS. Moreover, immunopanning can be performed in the standard culture vessel and is as simple as plating the cells on the adequate substrate for cell selection.

Materials 50 mM Tris·Cl, pH 9.5 Bovine serum albumin (BSA; Sigma, cat. no. A-3156) Anti-PSA-NCAM antibody (Iowa DSHB, http://dshb.biology.uiowa.edu/, cat. no. 5A5) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) 100 × 15–mm petri dish (bacterial grade, non-TC-treated; BD Falcon, cat. no. 351029) 1. Prepare the immunopanning cocktail:

50 mM Tris·Cl, pH 9.5 containing: 1% (w/v) BSA 50 μg/ml anti-PSA-NCAM antibody. 2. Coat the bottom surface of 100-mm non-tissue-culture-grade petri dishes by adding 4 to 5 ml per dish of the anti-immunopanning cocktail and incubating 30 min at 37◦ C. Flasks may also be coated by this protocol: use 2.5 ml to coat a 12.5-cm2 flask or 5 ml to coat a 75-cm2 flask.

3. Remove the cocktail. Wash petri dishes three times, each time with 5 ml PBS, then once with 5 ml PBS containing 1% BSA just before using. Do not allow the plates to dry. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

4. Cover extra plates (still containing the PBS/BSA) with foil and store at 4◦ C for up to 10 days.

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ASSESSING CELL VIABILITY Cell viability can be determined with the SYTOX blue nucleic acid stain (Molecular Probes/Invitrogen). Cells with compromised plasma membranes are labeled by SYTOX binding to nucleic acids and detected by fluorescence microscopy.

SUPPORT PROTOCOL 2

Materials Tris-buffered saline (TBS; see recipe) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) 1 μM SYTOX blue nucleic acid stain (Invitrogen Molecular Probes, cat. no. S7020) in PBS Microscope slides and coverslips Fluorescence microscope 1. Gently wash cells three times, each time with 5 ml TBS. 2. Harvest cells with a cell scraper and transfer them to a 15-ml tube. Resuspend in 2 ml PBS. Do not use enzymes.

3. Add 1 μl of 1 μM SYTOX to each tube (final concentration, 5 nM). 4. Incubate 12 min. 5. Remove the solution and wash the cells five times, each time with 5 ml TBS, centrifuging 3 min at 500 × g, room temperature, each time. 6. Resuspend the cells in 1 ml/tube of PBS. Take an aliquot and add a drop to a microscope slide. Add a coverslip and examine with a fluorescence microscope. 7. Determine the number of positive cells per random field in a fluorescence microscope and record as a percentage of the total number of cells in the field.

PROPAGATION OF RODENT NSCs AS TWO-DIMENSIONAL CULTURES NSCs can be propagated in two-dimensional (2-D) or three-dimensional (3-D) cultures. When attached, NSCs (2-D cultures) tend to grow faster and are therefore ideal for creating a large cell stock quickly before starting specific studies. In addition, we have developed a new method for expansion and maintenance of human NSC in NB-B-27 medium (see Reagents and Solutions) as multilayer adherent network (MAN) cultures, with increased proliferation rates compared to standard sphere-forming assays (UNIT 2D.3). In order to accommodate the difference in basal media, human NSC can either be initially derived in STMIIc media (replacing NB-B-27), or previously established hNSC cultures may be slowly transitioned away from the basal NB-B-27. Simply substitute 25% STMIIc for 1 week, followed by successive weeks at 50%, 75%, and finally 100% STMIIc after 1 month.

BASIC PROTOCOL 2

This procedure can be used every time cells are replated.

Materials Cultures of freshly isolated neural stem cells (Basic Protocol 1) or their progenitors Hanks’ buffered salt solution (HBSS) without Ca2+ or Mg2+ Complete stem cell medium (STMIIc; see recipe) Cell Freezing Medium, serum-free, 1× (Sigma, cat. no. C2639) Cell scraper 15-ml conical tubes

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Centrifuge (e.g., IEC Clinical) 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 12.5-cm2 and 75-cm2 tissue culture flasks (Falcon), anti-PSA-NCAM-coated (Support Protocol 1) 1.2-ml cryovials Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3) and freezing cells (Support Protocol 3) NOTE: All steps are performed at room temperature (20◦ C).

Collect the cells 1. When confluency of the neural stem cells has been reached, remove the supernatant conditioned medium (CM; save for step 4), add 5 ml of HBSS without Ca2+ or Mg2+ , detach the cells with a cell scraper, and transfer to a 15-ml conical tube (accommodating cells from one to three dishes). 2. Rinse the dish once with 2 ml HBSS (without Ca2+ or Mg2+ ), add it to the tube, and centrifuge 8 min at 450 × g. Discard the supernatant. 3. Resuspend the cell pellet in 3 ml STMIIc, dissociate gently using 18-G needle and syringe, and centrifuge 5 min at 450 × g. 4. Resuspend cells in 2 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM (use CM from step 1). Replate cells on anti-PSA-NCAM-coated plates as described in Basic Protocol 1. If you have repeated this process several times and the cell pellet is 0.5 ml volume or larger, divide the cell suspension into two parts. One part of the suspension will be used to start a frozen stock (see Support Protocol 3). The second half of the cell suspension is further dissociated using a needle (as described in Basic Protocol 1; however, the sieves (used in step 10 of Basic Protocol 1) are not necessary (single-cell suspension is obtained using needle and syringe), and replated as described below.

Replate the cells 5. Count the number of cells/ml and adjust the volume to 15 ml with a freshly prepared mixture of 2 parts STMIIc and 1 part CM (use CM from step 1). 6. Plate 2 ml of the cell suspension in each of five 12.5-cm2 cell culture flasks coated with anti-PSA-NCAM.

Passage the cells 7. Feed cells with a freshly prepared mixture of 2 parts STMIIc and 1 part CM every other day until they reach 80% to 90% confluency, and repeat steps 1 to 7 to increase the number of cells. 8. When four or more 12.5-cm2 flasks reach confluency, harvest the cells as described above, and seed the equivalent content of cells from three 12.5-cm2 flasks into one 75-cm2 flask coated with anti-PSA-NCAM. Feed the cells with a freshly prepared mixture of 2 parts STMIIc and 1 part CM in a total volume of 10 ml/flask. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

9. After 1 to 2 days, check to see if the culture medium is red. If the medium is turning orange, add 3 ml of STMIIc. Repeat this step as needed. Add 3 ml of STMIIc every day only if the medium changes color. If cells seem not to grow, but look healthy, or if the culture medium is not red but purple, you will need to remove 1/2 of the plating medium and bring the volume up to 10 ml with a freshly prepared mixture of 2 parts STMIIc and 1 part CM.

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If the opposite is true and the culture medium turns orange overnight, the cells have proliferated heavily, and you will need to replace the culture medium and seed more 75-cm2 flasks (one 75-cm2 flask per three 12.5-cm2 flasks).

10. Optional: When cells reach confluency, freeze the contents of one 75-cm2 flask (Support Protocol 3) in 1 ml of freezing medium in a cryovial. When propagating cells to create frozen stocks, we strongly recommend maintaining a “mother flask” by scraping most, but not all of the cells attached to the flask. After removing the detached cells, feed the mother flask with a 1:1 mixture of fresh STMIIc and CM to ensure continuity of these cultures (in case replated cells do not look healthy, grow slowly, or die).

11. Optional: After accumulating at least 10 to 15 vials of cryopreserved NSC in a frozen stock, grow NSCs as neurospheres (three-dimensional; see Alternate Protocol) for slower growth, allowing more time to devote to experiments. While cells are floating, even in STMIIc their metabolism seems slower, but if replated they behave normally.

FORMATION, PROPAGATION, AND MAINTENANCE OF NEUROSPHERES IN THREE-DIMENSIONAL CULTURES

ALTERNATE PROTOCOL

Suspension aggregate, or “neurosphere,” three-dimensional cultures are an alternative strategy to propagate NSCs at a slower pace than that of attached cells, while preserving most of the standard characteristics of a proper NSC. NSC suspension cultures are started from freshly dissociated NSC two-dimensional cultures (Basic Protocol 2) and grown in sterile Erlenmeyer flasks to prevent attachment and encourage free-floating spherical growth.

Materials Conditioned medium (see Basic Protocol 2) Complete stem cell medium (STMIIc; see recipe) Established NSC cultures (Basic Protocol 2) Glass Erlenmeyer flasks, 25-ml or 50-ml with cap 37◦ C incubator with shaker 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 50-ml conical tubes Centrifuge (e.g., IEC Clinical) 0.22-μm sterile filters NOTE: All steps are performed at room temperature (20◦ C). 1. After harvesting cells as described in Basic Protocol 2, resuspend the cells from one 75-cm2 flask in 6 ml fresh STMIIc. 2. Place 15 ml of fresh STMIIc and 3 ml of conditioned medium (CM) into a 25- or 50-ml Erlenmeyer flask (depending on number of cells). 3. Add 2 ml of the cell suspension (freshly harvested from two-dimensional cultures as described in Basic Protocol 2), and close the top of the flask partially to allow for O2 /CO2 exchange. Thus, one 75-cm2 flask will result in three Erlenmeyer flasks of neurospheres. Somatic Stem Cells

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4. Place the flask, continuously shaking at 90 rpm, in the incubator. If placing a shaker in the incubator is not an option due to safety regulations, the cell suspension may be placed directly into two noncoated petri dishes (bacterial grade) to prevent cell attachment.

5. Add 1.5 ml fresh STMIIc, every other day, and dissociate routinely (three times gently) with a syringe and needle in the same flask (as described for pellet dissociation in Basic Protocol 2) to keep the spheres at a small size. This process allows for increased sphere formation without the negative potential for spontaneous differentiation. It also allows cells more exposure to the fresh nutrients in the culture medium, helping preserve “stemness” in all cells.

6. When the culture medium turns orange overnight, collect the contents of the Erlenmeyer flask with a pipet and place into one 50-ml conical tube. Centrifuge for 6 min at 450 × g. 7. Slowly collect the CM (supernatant), filter (0.22-μm), and save for replating cells. 8. Resuspend the pellet in 4 ml of STMIIc to dissociate larger spheres. Add 21 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM. At this point, the neurospheres should all be easily dissociated. If some spheres remain large in spite of repeated dissociation, use the sieves (see Basic Protocol 1 materials list) to eliminate the large clumps, instead of vigorously dissociating them. This step will prevent significant cell death at the time of replating.

9. Seed cells on desired containers for experiments, or continue to propagate NSCs as two- or three-dimensional cultures (see Basic Protocol 2). SUPPORT PROTOCOL 3

CRYOPRESERVATION/THAWING OF NSC STOCKS We recommend collecting cells for frozen stocks at low passage number. Human NSC are cryopreserved using modified methods found elsewhere (Wakeman et al., 2009). In addition, the method formerly described for rat and mouse NSC (Espinosa-Jeffrey et al., 2002) can also be used to stock human NSC. The present methods are to be used for basic research and can be optimized for translational research if such cells are approved for the clinic. Therefore, we recommend the use of serum-free freezing medium as well as all other animal-free components as indicated within this unit. It would defeat the purpose to perform the full preparation and maintenance utilizing animal-free products and then expose the cells to fetal bovine serum or other types of sera at the moment of cryopreservation.

Materials NSC cultures ready for freezing (Basic Protocol 2) Hanks’ balanced salt solution (HBSS) without Ca2+ or Mg2+ Complete stem cell medium (STMIIc; see recipe) Cell Freezing Medium, serum-free, 1× (Sigma, cat. no. C2639) Liquid nitrogen Conditioned medium (CM; see Basic Protocol 2) Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Cell scrapers Centrifuge (e.g., IEC Clinical) 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 1.2-ml cryovials

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Cryogenic slow-freezing chamber (Nalgene, cat. no. EW-44400-00) 2-ml tubes (Fisher) Anti-PSA-NCAM coated tissue culture vessels (Support Protocol 1) Additional reagents and equipment for testing cell viability (Support Protocol 2) NOTE: All steps are performed at room temperature (20◦ C).

Collect and freeze the NSC 1. Allow NSCs to grow to 70% to 90% confluency. Remove all of the cell culture medium, add 5 ml of HBSS (without Ca2+ or Mg2+ ) to each 100-mm petri dish or 10 ml to each 75-cm2 flask, and detach cells by gently scraping the culturing surface. 2. Centrifuge the cells 8 min at 450 × g, and resuspend in 3 ml STMIIc medium. 3. Gently dissociate cells using an 18-G needle and syringe, centrifuge 8 min at 450 × g, and discard the supernatant. 4. Gently resuspend the pellet from one 100-mm petri dish or 75-cm2 flask in 1 ml of serum-free freezing medium. 5. Transfer the contents to a 1.2-ml cryovial, and place the vial(s) in a cryogenic freezer container overnight for slow freezing. 6. Next day, place the vials in liquid nitrogen for long-term storage.

Thaw NSCs 7. To reanimate NSCs, defrost cryovials quickly in a 37◦ C water bath, and transfer the contents of the vial to a 2-ml tube containing 1 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM at 37◦ C. 8. Centrifuge gently 5 to 7 min at 350 × g. 9. Remove the supernatant, add 1 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM at 37◦ C, resuspend the cell pellet, and remove a small aliquot to test the initial cell viability. 10. Count the number of viable cells in the tube (∼1 × 106 cells expected) as described in Support Protocol 2. 11. Plate cells onto an anti-PSA-NCAM-coated vessels (petri dishes or 75-cm2 flasks, plate the equivalent of 1 vial/75 cm2 flask). If the yield is lower, use 25-cm2 flasks to increase the cell density necessary for healthy growth. Seeding low-density cultures in large containers decreases the proliferation rate and might be detrimental to the culture.

12. To propagate NSCs after replating, proceed as described in Basic Protocol 2. Alternatively, when cells have reached 90% confluency, either freeze them or use them for experiments.

OLIGODENDROCYTE COMMITMENT IN TWO- AND THREE-DIMENSIONAL CULTURES

BASIC PROTOCOL 3

During development, the nutritional and environmental needs of cells change as they lose multipotency and become lineage restricted. The present system is based on the modification of nutrients contained in the cell culture medium and the percentage of CO2 needed to optimize and direct lineage restriction towards the oligodendrocyte (OL) phenotype. Like NSCs, OL progenitors (OLPs) can be propagated in two- and threedimensional cultures. When attached (two-dimensional cultures), OLPs grow faster and,

Somatic Stem Cells

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thus, two-dimensional cultures are ideal to create an OLP cell stock quickly before starting specific in vitro cell culture or in vivo transplantation studies. A diagram of the following steps can be found in Figure 2D.4.4.

Materials NSC cultures (2-D or 3-D; Basic Protocol 2 or Alternate Protocol) Hanks’ balanced salt solution (HBSS) without Ca2+ or Mg2+ OL specification medium (OSM-II; see recipe) Cell scraper 15-ml conical tubes 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) Anti-IgM coated 100-mm petri dishes or tissue culture flasks: prepare as in Support Protocol 1 but substitute goat anti-IgM antibody (ABR, sold by Thermo Scientific, cat. no. PA1-86106) for anti-PSA-NCAM antibody 37◦ C, 4.5% CO2 incubator 12-ml syringe (Tyco Healthcare, cat. no. 512852) Additional reagents and equipment for maintaining cells (Basic Protocol 1) NOTE: All steps are performed at room temperature (20◦ C). NOTE: We recommended precalibrating the percentage of CO2 1 day before plating the cells. If the incubator is shared with other people or needed at 5% for NSC propagation and maintenance, we recommend using tissue culture flasks for 2-D cultures instead of petri dishes. Close the cap of the flask completely and then open it one-quarter of a turn before placing in the incubator at 5% CO2 . For propagation and maintenance of OL spheres, the Erlenmeyer flask should also be kept open just enough to ensure O2 /CO2 exchange. When using 4.5% CO2 , loosen the caps of the flasks until half-way open. 1. When NSCs reach confluency, remove the supernatant (CM; save for use in subsequent steps), and add 5 ml of HBSS without Ca2+ or Mg2+ . 2. Detach the cells with a cell scraper, transfer into a 15-ml tube (which accommodates one to three petri dishes), rinse once with 2 ml of HBSS (without Ca2+ or Mg2+ ), and centrifuge 8 min at 450 × g. 3. Resuspend the cell pellet in 3 ml OSM-II medium and gently dissociate three times using a syringe and 18-G needle. Centrifuge 8 min at 450 × g to pellet the cells.

For two-dimensional OL cultures 4a. Resuspend the cells in a freshly prepared 1:1 mixture of OSM-II and CM. Seed cells on anti-IgM coated dishes or flasks. By using anti-IgM unconjugated antibody for coating, not only PSA-NCAM-positive cells will attach to the plate or dish, but also the cells that begin to express A2B5+ gangliosides. The panning strategy can be used at later stages to select OL cells at a single developmental stage (see Fig. 2D.4.1), e.g., pre-OL, which can be selected using anti-O4. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

The CM used here and in the following step is self-conditioned STMIIc.

5a. Maintain the cells as described in Basic Protocol 1 but using a mixture of 2 parts OSM-II and 1 part CM. From this point on, maintain the CO2 concentration in the incubator at 4.5%. Cells switch from NSCs to OLP (third stage shown in Fig. 2D.4.1) within 20 hr in contact with OSM, at which time the cells start to express transferrin (Tf).

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dissociation NSC in STM/OSM 2:1 2-D cultures

3-D cultures CM/OSM 2:1

overnight

slow propagation (oligospheres)

CM/OSM 1:1

fast propagation: plate on anti-PSA-NCAM

overnight

100% OSM

slow propagation

fast propagation

in vitro studies cryopreservation GDM

transplant studies

Figure 2D.4.4 Oligodendrocyte specification. The transition of NSC to commit to the OL lineage is brief, but sequential rather than abrupt. In order for cells to survive, they must acclimate to their new environment. OLP can be propagated to create frozen stocks as three-dimensional “oligosphere” cultures (shown on the left of the diagram) or frozen without propagation (as shown in the sequence in the center of the diagram). OLP can also be propagated in two-dimensional cultures for cryopreservation, for specific cell culture experiments, or for cell replacement therapies (as shown on the right side of the diagram).

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6a. Feed the cells with a mixture of 2 parts OSM-II and 1 part CM (i.e., self-conditioned OSM-II) every other day until they reach 80% to 90% confluency (3 to 5 days). This process can be repeated several times to attain a large number of cells for freezing (if desired).

For three-dimensional OL cultures 4b. Alternatively, to grow OL spheres to create/enrich a frozen stock of OLP, place the equivalent of 2 mm2 of cells (pellet size after cells are dissociated and in suspension) in a 25-ml Erlenmeyer flask with 15 ml of a mixture of 2 parts (10 ml) OSM-II and 1 part (5 ml) CM (i.e., self-conditioned OSM-II). If the pellet is 4 mm2 (∼12-15 × 106 OLP), use a 50-ml Erlenmeyer flask. Prepare the cell suspension and place in a total volume of 25 ml of a mixture of 2 parts OSM-II and 1 part CM (i.e., self-conditioned OSM).

5b. Feed OL spheres with fresh OSM-II every other day by adding 3 ml of freshly prepared OSM-II (no CM). 6b. When spheres start to become larger than 2 mm, gently dissociate by aspirating them one to two times in the same flask with the 18-G needle using a 12-ml syringe (sterile). 7. When the culture medium starts to turn orange, recover and centrifuge the spheres from one flask, and split cells into more Erlenmeyer flasks (1 to 4). These may be used for experiments or cryopreserved as previously described (Support Protocol 3). BASIC PROTOCOL 4

CULTURING OLIGODENDROCYTES FOR LINEAGE PROGRESSION AND MATURATION The nutritional needs for a committed cell within the OL lineage differ considerably as the cells progress and mature to the next developmental stage. These cells need to start synthesizing enzymes and proteins related to myelination; therefore, the energy demand is enormous compared to their earlier stage where migration and proliferation are the basic functions. The culture medium “GDM” (glial defined medium) was first designed to maintain 04+ , GC+/− , CNP+/− cells (for details. see “pre-OL” in Fig. 2D.4.1). Later, we realized that GDM also induced the transition of OLP to pre-OL (Espinosa de los Monteros et al., 1997). OL can be sustained at a given developmental stage by keeping them in one of the stage-specific culture media described here. Not all cell types offer the possibility for studying commitment and full differentiation when grown in culture, and one of the best examples comes from glial biology. Astrocytes grown in artificial cell culture conditions have allowed us to understand many of their functions and interactions with neurons and oligodendrocytes and how they play an integral part in mediating disease pathology; however, to our knowledge, there is no definitive proof of a fully matured, terminally differentiated astrocyte that can be studied throughout its progression and maturation in cell culture. In contrast to astroglial biology, we appear now to have the necessary tools to terminally differentiate oligodendrocytes in vitro, which even produce large amounts of compact myelin-like membranes in the absence of axons.

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Most cell culture methods for maintaining OL only allow the researcher to address commitment, survival, or maintenance in a cell type that will, by default, tend to progress from the progenitor stage into a more mature stage uncontrollably (i.e., beyond control by the researcher) when maintained in a fairly rich environment conducive to and promoting myelinogenic properties. However, in our model, the design of several culture media specific for multiple OL developmental stages provides us with the ability to control

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lineage progression of normal OL at multiple lineage transitions. In addition, these subtype-specific media allow us to determine potential deficiencies in diseased or stressed OL derived from transgenic/mutant animals and tissues donated by human subjects. Therefore, development of OL-lineage-specific media formulations allows us to further model disease mechanisms and determine how they affect OL at different stages of development. Furthermore, we can then use these data to design specialized culture media aimed at either further protecting the cell or preventing specific mechanisms from potentially occurring in OL-related diseases, or as part of their inherent injury response. Utilizing this platform, we can apply high-throughput small-molecule libraries to our defined media and determine how specialized formulations may effectively aid in the restoration and repair of degenerating tissue.

Materials OLPs (Basic Protocol 3, step 6a) OL specification medium (OSM-II; see recipe) GDM medium (see recipe) Recombinant human basic fibroblast growth factor (bFGF; Invitrogen) OLDEM medium (see recipe) Poly-D-lysine-coated wells/plates (see recipe) Additional reagents and equipment for oligodendrocyte differentiation in two-dimensional culture (Basic Protocol 3) Culture for pre-OLs 1. In order to obtain pre-OL (along the OL lineage), plate OLPs using OSM-II (as in Basic Protocol 3). As in previous steps, they may be propagated as OL spheres (three-dimensional) or as twodimensional cultures on anti-IgM coated flasks or petri dishes, or directly on cell-culture grade plastic. See Figure 2D.4.4 for options.

2. On the next day, remove one-half of the volume of the plating medium (OSM-II) and add the same volume of GDM. Continue incubation for a minimum of 2 days, or until 90% confluence is reached. 3. To obtain more OLP/pre-OL, grow cells as two- or three-dimensional cultures in the presence of bFGF (Fig. 2D.4.5). To keep progeny cells at the same stage as the parent cells, add 2 ml fresh GDM containing 20 ng/ml bFGF every other day until 90% confluence is reached. For cell replacement therapies, we suggest using cells at this stage (1 to 2 days after plating without bFGF; see alternatives described in Fig. 2D.4.4), as cells are still highly motile and readily migrate within the host post-natal and/or adult rodent brain and/or spinal cord.

4. To enhance maturation of cells into the next developmental stage, culture OL as two-dimensional cultures (Basic Protocol 3). Plate 1 × 105 cells/ml in GDM for at least 2 days (if plated in GDM without bFGF), or 4 days (if plated in GDM with bFGF) without further bFGF supplementation (Fig. 2D.4.5). After exposure to GDM, cells express myelin enzymes and proteins, and they display multipolar, branched cell processes, but not a myelin-like membrane. In addition, OL maintained in GDM for at least 4 days (without bFGF or any other factors) can be further induced to a fully mature myelinating stage. Also see Figure 2D.4.6.

5. To obtain fully mature OL, plate as two-dimensional cultures (Basic Protocol 3) at 1 × 105 cells/ml onto poly-D-lysine coated wells/plates or uncoated petri dishes in 10 ml of a 1:1 mixture of GDM and OLDEM (OL maturation medium). Culture for 1 to 5 days, then replace with 100% OLDEM for further culture (Fig. 2D.4.5).

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GDM bFGF 4 days

bFGF overnight

OLP propagation

maturation

bFGF overnight

bFGF overnight oligospheres OLDEM 1:1 1 to 5 days

GDM bFGF 2 days

bFGF overnight cocultures myelination studies

OLDEM 100% transplant studies

cryopreservation

mature myelinating

Figure 2D.4.5 Oligodendrocyte lineage progression and maturation. After commitment of NSC to the OL lineage, cells are propagated at the OLP stage to create a frozen stock (steps indicated on the left portion of the flow chart) or processed further for transplantation studies (as shown by the middle arrow on the diagram). To allow OLP to further mature along the OL lineage and become myelinated, cells are transitioned into OLDEM for at least 48 hr. Once OL have reached this stage of maturation, they are excellent for cell culture studies but are not recommended for cell grafting. Detachment from the substrate can damage the numerous delicate cell processes; therefore these cultures are no longer a quality source for cell transplants.

6. Every 4 days, feed the cells by replacing all of the culture medium with fresh OLDEM. Myelinating OL express myelin markers (see Fig. 2D.4.1, last two columns). The medium should look red, not orange. If it turns orange, add more medium while feeding the cells.

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

These cells will express myelin enzyme levels comparable to those found in pure myelin within 5 days after having been introduced to 100% OLDEM. As they mature, cells will synthesize myelin-like membranes in vitro even in the absence of neurons. They can be maintained for a number of weeks if they are kept subconfluent; however, if the culture becomes overcrowded, cells will deteriorate and die.

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B

C

OSM 2 days

OSM 3 days

GDM 1 day

DD

EE

F

Nestin Tf Tf

N estin T Nestin Tff

O 4 /MBP O4

G

H H

I

HuOLP, OSM 2 days

GDM 10 days, MBP

HuOL/Rat Neurons

A

Figure 2D.4.6 Phase-contrast view of neural stem cells derived from embryonic day 16 rat brain at passage number 2 (P2). NSC were plated and maintained in OSM for 2 days (A), 3 days (B), or 3 days in OSM then switched to GDM for 1 day (C). Cells in OSM still proliferate while in OSM. When cells from (A or B) are plated and maintained in OSM on poly-D-lysine-coated coverslips for 1 day, they start to display a bipolar or multipolar morphology (D) and most express the immature precursor marker, nestin (green) but not Tf (red), an early marker for OL. After 2 days in OSM, bipolar nestin+ cells coexpress transferrin (Tf) (E). After 4 days in OSM, cells were switched to GDM for 1 day; they developed numerous cell processes and coexpressed sulfatides (recognized by the anti-O4 antibody, green) and myelin basic protein (MBP; red) (F). Panels G to I are human cells. (G) Phase-contrast view of human NSCs (HFB-2050) acclimated and expanded in STM, then replated and maintained in OSM for 2 days. (H) OL derived from human NSCs (HFB-2050) were specified to the OL lineage with OSM and maintained in GDM for 10 days. OL matured and started to express MBP (red). (I) Rat cortical neurons (NFM-200-red) were cultured for 10 days, then human OLP derived from NSC (HFB-2050) were added in coculture for 24 hr. These cells were labeled with human nuclei marker (HuNu, green).

PROPAGATION OF OLIGODENDROCYTES FOR IN VITRO MYELINATION ASSAYS

BASIC PROTOCOL 5

To perform myelination studies in vitro, it is recommended to start with OL plated on plastic alone (rather than poly-D-lysine) and maintained in GDM for 2 days.

Materials OL plated on (uncoated) plastic tissue culture dishes (from Basic Protocol 4, step 2; also see Fig. 2D.4.5) GDM medium (see recipe) Conditioned medium (from GDM; Basic Protocol 4)

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OLDEM medium (see recipe) Cell scraper Neuronal cultures (Support Protocol 4) plated on poly-D-lysine-coated coverslips Complete Neurobasal-N medium for cortical neurons (see recipe) 40-μm cell strainers (BD Falcon, cat. no. 352340) NOTE: All steps are performed at room temperature (20◦ C). 1. Detach OL cells with cell scraper and centrifuge at 450 × g, in the original culture medium. 2. Remove the supernatant and resuspend the cells in CM plus fresh OLDEM (1:2). See Figure 2D.4.5 (right side).

3. To prepare a single-cell suspension, which is necessary for the next step, remove any cell clusters by passing the cell suspension through a 40-μm sieve and wash the sieve as described in Basic Protocol 1, step 10. 4. Count the cells (UNIT 1C.3) and adjust the cell suspension to ∼200,000 cells/ml in OLDEM medium. 5. At 10 days after plating, remove half the volume (250 μl) of culture medium from neuronal culture (Support Protocol 4) without disturbing the cells. The neurons for coculture can be either cortical neurons or dorsal root ganglion cells.

6. Slowly add 200 μl of the OL suspension (from step 4) to the wells of one 24-well plate containing the neuronal cultures. To complete the original total volume in each well, add 50 μl/well of complete Neurobasal N medium for cortical neurons. 7. Follow the cocultures for at least 10 days. To feed, on day 5 after starting coculture, remove one half of the CM from each well (250 μl) and replace with fresh OLDEM. Repeat OLDEM feeding once a week. There is no need to reapply Neurobasal N medium. If the cultures are not overcrowded, they can be kept for at least 4 weeks. SUPPORT PROTOCOL 4

PREPARATION OF CORTICAL NEURONS Cortical neurons are one cell type that is used for coculture with the OLs to assess myelination.

Materials Complete Neurobasal-N medium for cortical neurons (see recipe) Poly-D-lysine-coated (see recipe) coverslips in wells of 12- or 24-well plates 37◦ C 4.5% CO2 incubator, 95% humidified Combustion Test Kit (Bacharach, cat. no. 10-500; http://www.bacharach-inc.com) Additional reagents and materials for isolation of rodent brain cells (see Basic Protocol 1) NOTE: All steps are performed at room temperature (20◦ C).

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

1. Prepare and dissociate embryonic rat brain tissue as described in Basic Protocol 1, steps 1 to 8, except use complete Neurobasal-N medium in step 5 of that protocol (instead of STMIIc) and dissect the brains in Neurobasal-N medium. 2. Add 2 to 4 ml of Neurobasal-N medium to the chunks left over in the dissociation tube and dissociate again five to eight times. 3. Filter the suspension of dissociated cells through 230-μm and 140-μm sieves to remove cell clusters.

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4. Rinse the two sieves sequentially with Neurobasal-N containing 1% BSA at room temperature as described in Basic Protocol 1, step 10, and add this medium to the tubes containing the cells. 5. Collect the cells by centrifugation in the culture tubes 8 min at 400 × g. 6. Discard the supernatant, very gently as the pellet is very loose. 7. Resuspend the pellet in 4 ml of fresh Neurobasal-N medium with a 5-ml pipet by gently triturating (i.e., pipetting up and down) two or three times. Bring the volume to 12 ml (or the equivalent of 1 embryo/ml) with 2 parts of fresh Neurobasal N medium and 1 part of conditioned medium. 8. Assess cell viability with SYTOX (Support Protocol 2), count cells using a hemacytometer (UNIT 1C.3), and plate onto poly-D-lysine coated coverslips inside wells of 12or 24-well plates at 2 × 105 cells/well in 700 μl complete Neurobasal N medium for cortical neurons. 9. Incubate plated cells at 37◦ C with 4.5% CO2 /95% humidity and monitor CO2 with a Combustion Test Kit because most electronic panels do not give an accurate reading. 10. Every third day, add 50 μl/well complete Neurobasal-N complete medium. On the sixth day after plating, remove one-quarter of the medium and add the corresponding volume of fresh complete Neurobasal-N medium. Neurons are ready for coculture (Basic Protocol 5) 10 days after plating.

TRANSPLANTATION OF OL PROGENITORS INTO NEONATAL RATS Neural progenitor cells and their differentiated OL counterparts can be stereotaxically transplanted into the newborn developing rat brain relatively noninvasively as previously described (Snyder et al., 1997; Flax et al., 1998; Espinosa-Jeffrey et al., 2002). Similar results can be obtained with variations on the transplant method that are more suitable depending on the needs of the host brain and the type of study (Yandava et al., 1999; Ourednik et al., 2001, 2002; Park et al., 2002a; Teng et al., 2002; Wakeman et al., 2006; Lee et al., 2007; Redmond et al., 2007). A selection of detailed protocols for neonatal and adult mouse transplantation are described elsewhere (Espinosa de los Monteros et al., 1992, 1993a,b; Yan et al., 2004; Lee et al., 2008; Wakeman et al., 2009; UNIT 2D.3). Upon implantation into the lateral ventricles, donor cells engraft and migrate from the subventricular zone into the host corpus callosum, caudate putamen, and rostral migratory stream (RMS) in much the same manner as host NSC.

BASIC PROTOCOL 6

Materials Neonatal rat pup, post-natal day 0 to 5 (P0 to P5) 70% ethanol Dulbecco’s phosphate-buffered saline (DPBS; without calcium or magnesium, e.g., Cellgro, cat. no. 21-031-CV), sterile Microcentrifuge tube with cell sample (suspended in PBS; may be from various protocols in this unit depending on experimental question to be addressed) Borosilicate glass (Sutter Instrument Co., cat. no. B100–75-15) Micropipet puller (Sutter Instrument Co., Model P-87) Aspirator tube assemblies for calibrated microcapillary pipets (Sigma-Aldrich, cat. no. A5177–5EA) Fiber-optic light source for transillumination Warming pad Warm-water glove balloon

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Additional reagents and equipment for preparing injection micropipet (Lee et al., 2008) NOTE: Required materials may vary depending upon the grafting method of choice. 1. Prepare calibrated drawn borosilicate glass micropipet using borosilicate glass and a micropipet puller (Lee et al., 2008). 2. Anesthetize the neonatal rat pup by placing the pup on wet ice for ∼1.5 to 3 min until the animal no longer retains locomotion or responds to gentle toe and tail pinch. Carefully monitor the pup and immediately proceed to transplantation.

3. Insert a calibrated, drawn borosilicate glass micropipet into an aspirator tube assembly. Just prior to drawing up the cells, rinse the micropipet by drawing up and then expelling 5 μl of 70% ethanol five times, followed by sterile DPBS ten times to clean the needle.

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Figure 2D.4.7 Human OL derived from (HFB-2050) human fetal NSC were labeled with fluorescent fast blue (FB; Sigma, cat. no. F-5756). A total of 60,000 cells were grafted into the corpus callosum (CC) of P(5) rat pups born to a myelin-deficient (md) carrier mother. At a time point 23 days after grafting, samples were harvested and examined. Grafted NSC survived and migrated extensively within the host brain parenchyma extending along the corpus callosum (CC) and caudate putamen (CPu). In the sketch, dots represent the location where FB+ cells were found. The sketch represents a sagittal view of the transplanted rat brain at 28 days of age, IS indicates where cells were implanted.

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4. Gently flick sample in microcentrifuge tube prior to filling the needle, wipe the tube with 70% ethanol, and uncap the tube. 5. Slowly draw 4 to 5 μl cell suspension into the micropipet. 6. Loosely secure the head of the anesthetized pup and place directly over the light source to visualize the eyes and bregma. 7. Carefully insert the glass needle into the head at the midline between eye and bregma and slowly inject 2 to 5 μl cell suspension at 5 × 104 cells/μl into the lateral ventricle of either the left or the right hemisphere. Slowly remove the needle and check for leakage through the needle track. Repeat step 6 into the contralateral hemisphere. In addition to the lateral ventricles, NSC can also be transplanted into the striatum, the substantia nigra (SN), and corpus callosum (CC; Bjugstad et al., 2005, 2008; Redmond et al., 2007). Upon implantation into the CC of the host, HFB-2050 donor cells recognized by the fluorescent Fast Blue (FB) label migrated along the CC and into the caudate putamen (Fig. 2D.4.7). Precommitted OL can also be placed locally within focal sites of injury to decrease the need for extensive migration.

8. After the injection, warm the pup by placing on a warm-water glove balloon or heating pad to increase the body temperature before returning to the mother.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Basal stem cell medium (STM-II) Prepare 1 liter DMEM (low glucose, without glutamine, with sodium pyruvate; Invitrogen, cat. no. 11995-065). Supplement with the following:

5 mg insulin (Sigma, cat. no. I-5500) 50 mg transferrin (Sigma, cat. no. T-2252) 16.1 mg putrescine (Sigma, cat. no. P-7505) 20 nM (6.29 mg/liter) progesterone (Sigma, cat. no. P-7556) 8 μg sodium selenite (Sigma, cat no. S-5261); add 10 μl/liter of 0.8 mg/ml stock solution in PBS 2.2 g sodium bicarbonate (Fisher, cat. no. S233-500) 1 ml 10,000 U/ml penicillin/10 mg/ml streptomycin (Sigma, cat. no. P-4333) 1 ml 50 mg/ml kanamycin (Sigma, cat. no. K-0254) Store up to 2 weeks at 4◦ C This medium is used for rat and human NSC. STM-II is a variation of the original STM medium we previously described (Espinosa et al., 2002; UCLA case number 2002-475, formula available by Materials Transfer Agreement). STM-II yields results comparable to those obtained with STM.

Complete stem cell medium II (STMIIc) Just before use, combine 500 ml basal stem cell medium (STM-II; see recipe) and 500 ml NB-B27 medium (see recipe). This medium is used for plating, maintenance, and propagation of NSCs.

OL specification medium (OSM-II) Mix freshly prepared complete stem cell medium (STMIIc; see recipe) with freshly prepared glia defined medium (GDM; see recipe) at a 1:1 (v/v) ratio. This medium was formerly named OTM (Espinosa-Jeffrey et al., 2002); OSM-II derives from STM-II (see recipe).

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Glia defined medium (GDM) Combine 1 liter double-distilled water and one package DMEM/F12 medium (high glucose), then supplement with:

5 mg insulin (Sigma, cat. no. I-5500) 50 mg transferrin (Sigma, cat. no. T-2252) 16.1 mg putrescine (Sigma, cat. no. P-7505)) 2.2 g sodium bicarbonate (Fisher, cat no. S233-500) 4.6 g D-(+)-galactose (Sigma, cat. no. G-0625) 8 μg sodium selenite (Sigma, cat. no. S-5261): prepare 0.8 mg/ml stock solution in PBS (Sigma, cat. no. P-5368) and add 10 μl of this stock per liter medium 1 ml 50 mg/ml kanamycin (Sigma, cat. no. K-0254) Filter-sterilize through a 0.22-μm filter Prepare fresh From Espinosa de los Monteros et al. (1988, 1997)

Neurobasal-B-27 (NB-B-27) human neural stem cell proliferation medium Prepare 484 ml Neurobasal medium without Normocin, heparin, vitamin A, or LIF (Invitrogen, cat. no. 21103-049). Store up to 2 weeks at 4◦ C. Just before use in preparing STMIIc medium (see recipe), supplement with:

10 ml B-27 supplement without vitamin A (Invitrogen, cat. no. 12587-010) 5 ml GlutaMAX (Invitrogen, cat. no. 35050-061) 8 μg/ml heparin (Sigma, cat. no. H-3149) 2 ng/ml basic fibroblast growth factor (bFGF; Invitrogen, cat. no. 13256-029) 10 ng/ml leukemia inhibitory factor (LIF; Millipore, cat. no. LIF-1010) Neurobasal-N medium for cortical neurons, complete 1 liter Neurobasal medium (Invitrogen, cat. no. 21103-049) supplemented with: 5 mg insulin (Sigma, cat. no. I-5500)) 50 mg transferrin (Sigma, cat. no. T-2252) 8 μg sodium selenite (Sigma, cat. no. S-5261): prepare 0.8 mg/ml stock solution in PBS (Sigma, cat. no. P-5368) and add 10 μl of this stock per liter medium 2.2 g sodium bicarbonate (Fisher, cat. no. S233-500) 1 ml kanamycin (Sigma, cat. no. K-0254) Just before using, add the following to the supplemented Neurobasal-N to make the complete medium: 1:50 (v/v) B-27 supplement with vitamin A (Invitrogen, cat. no. 17504-044) 20 ng/ml recombinant basic bFGF (Invitrogen, cat. no. 13256-029) OLDEM medium Prepare glia defined medium (GDM; see recipe), but omit transferrin.

Poly-D-lysine coated wells/plates/coverslips

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Prepare a stock solution by dissolving 100 mg poly-D-lysine in 100 ml water and filter sterilize through a 0.22-μm filter. Store in 5-ml aliquots at –20◦ C. When ready to use, dilute 1 part stock solution with 9 parts water to prepare 100 μg/ml working solution. Fill tissue culture dishes or wells with the working solution (and/or place coverslips to be coated into wells of 12- or 24-well plate) and incubate 1 hr in a humidified 37◦ C, 5% CO2 incubator, then remove solution by vacuum aspiration and allow surface to dry. Store coated tissue culture ware up to 3 months at 4◦ C. Use diluted solutions only once, but unused diluted aliquots can be stored up to 3 months at 4◦ C.

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Tris-buffered saline (TBS) 2.42 g/liter Tris base 29.22 g/liter NaCl Adjust pH to 7.5 with HCl Store at room temperature up to 1 year, under sterile conditions COMMENTARY Background Information The described culturing system allows for the production of relatively homogeneous primary OL cultures in adequate numbers for cryopreservation. These cell stocks can be used for basic research in further in vitro studies. Moreover, these cells are never exposed to animal or human sera, and therefore remain as suitable candidates for cell replacement therapies in developmental disorders of the central nervous system (CNS) as well as neurodegenerative diseases. Numerous methods and culture media described in the literature (even before, the times of NSC) were the basis for the optimization of the culture media formulations described here (some examples are Botenstein and Sato, 1979; Saneto and de Vellis, 1985; Espinosa de los Monteros et al., 1988; Yang et al., 2005; UNIT 2D.1). Undoubtedly, all previous reports on how to obtain and culture OL derived from NSC have also been instrumental in designing the present protocols. For example, the group of Lachapelle and Baron-Van Evercooren described floating oligospheres derived from newborn rat brain (AvellanaAdalid et al., 1996). This concept has been applied to NSC to generate OLs by Zhang et al. (1998) and Espinosa-Jeffrey et al. (2002) and in the protocols in this unit. Zhang et al. (1998) described the use of B104 neuroblastoma cell–conditioned medium (B104CM) to induce the oligodendrocyte phenotype on neurospheres and induce proliferation. This approach provides OL for many kinds of studies, but they are unsuitable as donor cells for cell-replacement therapies to be used in translational studies, because they are produced using uncharacterized conditioned medium from B104 cells that have been grown in the presence of fetal bovine serum (as originally described by Louis et al., 1992). An example of the use of the protocols described can be found in Chattopadhyay et al., (2008).

Critical Parameters We want to emphasize that fate restriction towards commitment from NSC to OLP (as defined in Basic Protocol 3) becomes irreversible

after NSCs have been in OSM for at least 20 hr in either two- or three-dimensional cultures. Therefore, the progeny of these cells will define a homogeneous OLP population, ideal for biochemical, toxicological, and pharmacological studies, and also serve as an appropriate and reproducible source of committed cells to be used in cell therapy studies. Phenotype reversal of induced OLPs may be possible with genetic manipulation, but we have not attempted such studies to date. Always monitor the concentration of CO2 with a Combustion Test Kit, as most electronic panels do not provide an accurate reading. The proper lineage progression relies on precise control of CO2 to maintain a pH that should remain accurate and controlled.

Troubleshooting Human NSC are more fragile than their rodent counterparts; therefore, we recommend dissociation protocols that favor as little mechanical stress as possible. In our hands, enzymatic dissociation with 2 to 4 ml Accutase (Millipore) at 37◦ C for 3 to 5 min or light mechanical trituration through an 18-G needle (three to five times) is sufficient to dissociate hNSC into single cells and small (2- to 6-cell) clusters. Detailed methodology can be found elsewhere (Wakeman et al., 2009; UNIT 2D.3).

Anticipated Results OLPs obtained utilizing this system are plated on anti-PSA/NCAM plates and will attain a bipolar morphology if maintained in freshly supplemented OSM. Cells can also be plated directly onto plastic (tissue-culture grade). The morphology in that case may look more flattened or fibroblastic, but if maintained in fresh OSM, the early markers such as Olig2, Tf, PDGF-R, and NG2 will be expressed. At this stage, cells are still highly motile but will migrate less if plated onto poly-D-lysine. During this time, cells attain a more mature phenotype that truly represents their in vivo counterparts. Our culture media formulation includes the minimum and sufficient nutrients to support a given developmental stage; thus, cells cannot

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be kept indefinitely under these conditions because the substratum dictates the organization of the molecules on the cell membrane and poly-D-lysine confers a more permanent adhesion to the cells. Consequently, they would have the tendency to mature based on the signals coming from the cell membrane– substrate interaction (Linnemann and Bock, 1989; Mauro et al., 1994). Unfortunately, cells will not survive or remain healthy if maintained in unreplenished OSM as 2-D cultures, due to a lack of nutrients to support their transition to the next developmental stage. These cells will survive well if fed with OSM to renew the growth factors. The same concept applies to the transition to more mature OL stages. The nutrients and substrate together contribute to support cell signaling that will result in the formation of multiple cell processes followed by the synthesis of myelin components and their organization for membrane formation.

Time Considerations

Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

The initial dissection and preparation of the primary cell suspension takes ∼2 hr. From the moment cells are plated on anti-PSA-NCAM (if fed regularly with fresh humoral factors), 100-mm dishes can be confluent within 3 to 4 days. Thus, generating 20 vials of rat NSCs for cryostorage would take ∼16 days. The generation of OLP from rNSC takes ∼24 hr; however, generating OLP in high numbers (15 vials) for storage would take 4 to 6 weeks. Lineage progression of rat OL towards more mature phenotypes takes ∼48 hr in the specific culture medium (GDM or OLDEM). In addition, OL will still proliferate in GDM, but at a much slower rate. Both GDM and OLDEM media are favorable to protein synthesis but less favorable for cell proliferation. Previously isolated ES cells and their NSC derivatives will need a longer period of time to provide high numbers of NSC for frozen stocks. This time will vary depending on the origin of the sample. We have had similar success directing NSC from several species, utilizing the same chemically defined media; however, incubation times may need to be increased for full maturation in higher-order mammals, such as primates. Induced cells lose NSC characteristics and acquire OLP features within 72 hr, yet their cell cycle is much slower, and, therefore, it would be necessary to propagate these cells 8 to 10 weeks to be able to create a healthy stock (six to eight vials) of human OLP. Previously established NSC lines (Snyder et al., 1992) can also be propagated

and specified into the OL phenotype using the system described here.

Acknowledgements A.E. and J. de V. thank the MRRC Media Core for preparation of figures and Dr. D. Birt for photograph of the dissection set up. This work (J.de V. and A.E.) was supported in part by PPGHD065-76 and by a Pilot grant from the National Multiple Sclerosis Society PP1498. D.R.W. thanks M. Hudson for critical review and comments. D.R.W. is supported in part by the American Parkinson’s Disease Association, HHMI Med-Into-Grad Training Fellowship, and the UCSD-NIH Training Fellowship in Clinical Genetics.

Conflict of Interest Statement The authors acknowledge no conflict of interest.

Literature Cited Ahn, S.M., Byun, K., Kim, D., Lee, K., Yoo, J.S., Kim, S.U., Jho, E.H., Simpson, R.J., and Lee, B. 2008. Olig2-induced neural stem cell differentiation involves downregulation of Wnt signaling and induction of Dickkopf-1 expression. PLoS ONE 3:e3917. Arenander, A.T. and De Vellis, J. 1995. Earlyresponse gene expression in glial cells. In Neuroglia (H. Kettenmann and B.R. Ransom, eds.) pp. 510-522. Oxford University Press, New York. Avellana-Adalid, V., Nait-Oumesmar, B., Lachapelle, F., and Baron-Van Evercooren, A. 1996. Expansion of rat oligodendrocyte progenitors into proliferative “oligospheres” that retain differentiation potential. J. Neurosci. Res. 45:558-570. Bjugstad, K.B., Redmond, D.E. Jr., Teng, Y.D., Elsworth, J.D., Roth, R.H., Blanchard, B.C., Snyder, E.Y., and Sladek, J.R. Jr. 2005. Neural stem cells implanted into MPTP-treated monkeys increase the size of endogenous tyrosine hydroxylase-positive cells found in the striatum: A return to control measures. Cell Transplant. 14:183-192. Bjugstad, K.B., Teng, Y.D., Redmond, D.E. Jr., Elsworth, J.D., Roth, R.H., Cornelius, S.K., Snyder, E.Y., and Sladek, J.R. Jr. 2008. Human neural stem cells migrate along the nigrostriatal pathway in a primate model of Parkinson’s disease. Exp. Neurol. 211:362-369. Bottenstein, J.E. and Sato, G.H. 1979. Growth of a rat neuroblastoma cell line in serum-free supplemented medium. Proc. Natl. Acad. Sci. U.S.A. 76:514-517. Chattopadhyay, N., Espinosa-Jeffrey, A., TfeltHansen, J., Yano, S., Bandyopadhyay, S., Brown, E.M., and de Vellis, J. 2008. Calcium receptor expression and function in oligodendrocyte commitment and lineage progression: Potential impact on reduced myelin basic

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protein in CaR-null mice. J. Neurosci. Res. 86:2159-2167. De Filippis, L., Lamorte, G., Snyder, E.Y., Malgaroli, A., and Vescovi, A.L. 2007. A novel, immortal, and multipotent human neural stem cell line generating functional neurons and oligodendrocytes. Stem Cells 25:2312-2321. Espinosa de los Monteros, A., Roussel, G, Neskovic, N.M. and Nussbaum, J.L. 1988. A chemically defined medium for the culture of mature oligodendrocytes. J. Neurosci. Res. 19:202-211. Espinosa de los Monteros, A., Zhang, M., Gordon, M., Aymie, M., and de Vellis, J. 1992. Transplantation of cultured premyelinating oligodendrocytes into normal and myelin-deficient rat brain. Dev. Neurosci. 14:98-104. Espinosa de los Monteros, A., Zhang, M.-S., and de Vellis, J. 1993a. O2A progenitor cells transplanted into the neonatal rat brain develop Into oligodendrocytes but not astrocytes. Proc. Natl. Acad. Sci. U.S.A. 90:50-54. Espinosa de los Monteros, A., Bernard, R., Tiller, B., Rouget, P., and de Vellis, J. 1993b. Grafting of fast blue labeled glial cells into neonatal rat brain: Differential survival and migration among cell types. Int. J. Dev. Neurosci. 11:625-639. Espinosa de los Monteros, A. and de Vellis, J. 1996. Vulnerability of oligodendrocytes to environmental insults: Potential for recovery. In The Role of Glia in Neurotoxicity (M. Aschner and H.K. Kimelberg, eds.) pp. 15-46. CRC Press, Boca Raton, Fla. Espinosa de los Monteros, A., Yuan, J., McCartney, D., Madrid, B.R., Cole, R., Kanfer, J.N., and de Vellis, J. 1997. Acceleration of the maturation of oligodendroblasts into oligodendrocytes and enhancement of their myelinogenic properties by a chemically defined medium. Dev. Neurosci. 19:297-311. Espinosa-Jeffrey, A., Becker-Catania, S., Zhao, P.M., Cole, R., and de Vellis, J. 2002. Phenotype specification and development of oligodendrocytes and neurons from rat stem cell cultures using two chemically defined media. J. Neurosci. Res. 69:810-825. Flax, J.D., Aurora, S., Yang, C., Simonin, C., Wills, A.M., Billinghurst, L.L., Jendoubi, M., Sidman, R.L., Wolfe, J.H., Kim, S.U., and Snyder, E.Y. 1998. Engraftable human neural stem cells respond to developmental cues, replace neurons, and express foreign genes. Nat. Biotechnol. 16:1033-1039. Kim, H.T., Kim, I.S., Lee, I.S., Lee, J.P., Snyder, E.Y., and Park, K.I. 2006. Human neurospheres derived from the fetal central nervous system are regionally and temporally specified but are not committed. Exp. Neurol. 199:222-235.

of immortal human neural stem cell line with multipotent differentiation property. In Methods in Molecular Biology, Vol. 438: Neural Stem Cells, 2nd ed. (L.P. Weiner, ed.) pp.103-121. Humana Press, Totowa, N.J. Larsen, E.C., Kondo, Y., Fahrenholtz, C.D., and Duncan, I.D. 2008. Generation of cultured oligodendrocyte progenitor cells from rat neonatal brains. Curr. Protoc. Stem Cell Biol. 6:2D.1.1-2D.1.13. Lee, H.J., Kim, K.S., Kim, E.J., Choi, H.B., Lee, K.H., Park, I.H., Ko, Y., Jeong, S.W., and Kim, S.U. 2007. Brain transplantation of immortalized human neural stem cells promotes functional recovery in mouse intracerebral hemorrhage stroke model. Stem Cells 25:12041212. Lee, J.P., McKercher, S., M¨uller, F.J., and Snyder, E.Y. 2008. Neural stem cell transplantation in mouse brain. Curr. Protoc. Neurosci. 42:3.10.13.10.23. Linnemann, D. and Bock, E. 1989. Cell adhesion molecules in neural development. Dev. Neurosci. 11:149-173. Louis, J.C., Magal, E., Muir, D., Manthorpe, M., and Varon, S. 1992. CG4, a new bipotential glial cell line from rat brain, is capable of differentiating in vitro either mature oligodendrocytes or type-2 astrocytes. J. Neurosci. Res. 31:193-204. Mauro, V.P., Wood, I.C., Krushel, C., Crossin, K.L., and Edelman, G.M. 1994. Cell adhesion alters gene transcription in chicken embryo brain cells and mouse embryonal carcinoma cells. Proc. Natl. Acad. Sci. U.S.A. 91:2868-2872. M¨uller, F.J., Snyder, E.Y., and Loring, J.F. 2006. Gene therapy: Can neural stem cells deliver? Nat. Rev. Neurosci. 7:75-84. Neman, J. and De Vellis, J., eds. 2008. Handbook of Neurochemistry and Molecular Neurobiology: Myelinating Cells in the Central Nervous System—Development, Aging, and Disease. Springer, New York. Ourednik, V., Ourednik, J., Flax, J.D., Zawada, W.M., Hutt, C., Yang, C., Park, K.I., Kim, S.U., Sidman, R.L., Freed, C.R., and Snyder, E.Y. 2001. Segregation of human neural stem cells in the developing primate forebrain. Science 293:1820-1824. Ourednik, J., Ourednik, V., Lynch, W.P., Schachner, M., and Snyder, E.Y. 2002. Neural stem cells display an inherent mechanism for rescuing dysfunctional neurons. Nat. Biotechnol. 20:11031110. Palmer, T.D., Schwartz, P.H., Taupin, P., Kaspar, B., Stein, S.A., and Gage, F.H. 2001. Cell culture: Progenitor cells from human brain after death. Nature 411:42-43.

Kim, S.U. 2004. Human neural stem cells genetically modified for brain repair in neurological disorders. Neuropathology 24:159-171.

Park, K.I., Teng, Y.D., and Snyder, E.Y. 2002a. The injured brain interacts reciprocally with neural stem cells supported by scaffolds to reconstitute lost tissue. Nat. Biotechnol. 20:1111-1117.

Kim, S.U., Nagai, A., Nakagawa, E., Choi, H.B., Bang, J.H., Lee, H.J., Lee, M.A., Lee, Y.B., and Park, I.H. 2008. Production and characterization

Park, K.I., Ourednik, J., Ourednik, V., Taylor, R.M., Aboody, K.S., Auguste, K.I., Lachyankar, M.B., Redmond, D.E., and Snyder, E.Y. 2002b. Global

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gene and cell replacement strategies via stem cells. Gene Ther. 9:613-624. Redmond, D.E. Jr., Bjugstad, K.B., Teng, Y.D., Ourednik, V., Ourednik, J., Wakeman, D.R., Parsons, X.H., Gonzalez, R., Blanchard, B.C., Kim, S.U., Gu, Z., Lipton, S.A., Markakis, E.A., Roth, R.H., Elsworth, J.D., Sladek, J.R. Jr., Sidman, R.L., and Snyder, E.Y. 2007. Behavioral improvement in a primate Parkinson’s model is associated with multiple homeostatic effects of human neural stem cells. Proc. Natl. Acad. Sci. U.S.A. 104:12175-12180. Saneto, R.P. and de Vellis, J. 1985. Characterization of cultured rat oligodendrocytes proliferating in a serum-free chemically defined medium. Proc. Natl. Acad. Sci. U.S.A. 82:3509-3513. Schwartz, P.H., Bryant, P.J., Fuja, T.J., Su, H., O’Dowd, D.K., and Klassen, H. 2003. Isolation and characterization of neural progenitor cells from post-mortem human cortex. J. Neurosci. Res. 74:838-851. Snyder, E.Y., Deitcher, D.L., Walsh, C., ArnoldAldea, S., Hartwieg, E.A., and Cepko, C.L. 1992. Multipotent neural cell lines can engraft and participate in development of mouse cerebellum. Cell 68:33-51. Snyder, E.Y., Yoon, C., Flax, J.D., and Macklis, J.D. 1997. Multipotent neural precursors can differentiate toward replacement of neurons undergoing targeted apoptotic degeneration in adult mouse neocortex. Proc. Natl. Acad. Sci. U.S.A. 94:11663-11668. Svendsen, C.N., Caldwell, M.A., and Ostenfeld, T. 1999. Human neural stem cells: Isolation, expansion and transplantation. Brain Pathol. 9:499-513. Teng, Y.D., Lavik, E.B., Qu, X., Park, K.I., Ourednik, J., Zurakowski, D., Langer, R., and Snyder, E.Y. 2002. Functional recovery following traumatic spinal cord injury mediated by a unique polymer scaffold seeded with neural stem cells. Proc. Natl. Acad. Sci. U.S.A. 99:3024-3029.

Villa, A., Snyder, E.Y., Vescovi, A., and Mart´ınezSerrano, A. 2000. Establishment and properties of a growth factor-dependent, perpetual neural stem cell line from the human CNS. Exp. Neurol. 161:67-84. Wakeman, D.R., Crain, A.C., and Snyder, E.Y. 2006. Large animal models are critical for rationally advancing regenerative therapies. Regenerative Med. 1:405-413. Wakeman, D.R., Hofmann, M.R., Teng, Y.D., and Snyder, E.Y. 2009. Derivation, expansion, and characterization of human fetal forebrain neural stem cells. In Human Cell Culture: Adult Stem Cells: Vol. 7. (J.R. Masters and B.O. Palsson, eds.). Springer, Dordrecht, The Netherlands. Williams, W.C. 2nd and Gard, A.L. 1997. In vitro death of jimpy oligodendrocytes: Correlation with onset of DM-20/PLP expression and resistance to oligodendrogliotrophic factors. J. Neurosci. Res. 50:177-189. Wysocki, L.J. and Sato, V.L. 1978. Panning for lymphocytes: A method for cell selection. Proc. Natl. Acad. Sci. U.S.A. 75:2844-2848. Yan, J., Welsh, A.M., Bora, S.H., Snyder, E.Y., and Koliatsos, V.E. 2004. Differentiation and tropic/trophic effects of exogenous neural precursors in the adult spinal cord. J. Comp. Neurol. 480:101-114. Yandava, B.D., Billinghurst, L.L., and Snyder, E.Y. 1999. Global cell replacement is feasible via neural stem cell transplantation: Evidence from the dysmyelinated shiverer mouse brain. Proc. Natl. Acad. Sci. U.S.A. 96:7029-7034. Yang, Z., Watanabe, M., and Nishiyama, A. 2005. Optimization of oligodendrocyte progenitor cell culture method for enhanced survival. J. Neurosci. Methods 149:50-56. Zhang, S.C., Lundberg, C., Lipitz, D., O’Connor, L.T., and Duncan, I.D. 1998. Generation of oligodendroglial progenitors from neural stem cells. J. Neurocytol. 27:475-489.

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Isolation and Culture of Ventral Mesencephalic Precursor Cells and Dopaminergic Neurons from Rodent Brains

UNIT 2D.5

Jan Pruszak,1,2 Lothar Just,3 Ole Isacson,2 and Guido Nikkhah1 1

Freiburg University Hospital, Freiburg, Germany Harvard Medical School, McLean Hospital, Belmont, Massachusetts 3 Institute of Anatomy, Center for Regenerative Biology and Medicine, Eberhardt-Karls-University T¨ubingen, T¨ubingen, Germany 2

ABSTRACT The ability to isolate ventral midbrain (VM) precursor cells and neurons provides a powerful means to characterize their differentiation properties and to study their potential for restoring dopamine (DA) neurons degenerated in Parkinson’s disease (PD). Preparation and maintenance of DA VM in primary culture involves a number of critical steps to yield healthy cells and appropriate data. Here, we offer a detailed description of protocols to consistently prepare VM DA cultures from rat and mouse embryonic fetal-stage midbrain. We also present methods for organotypic culture of midbrain tissue, for differentiation as aggregate cultures, and for adherent culture systems of DA differentiation and maturation, followed by a synopsis of relevant analytical read-out options. Isolation and culture of rodent VM precursor cells and DA neurons can be exploited for studies of DA lineage development, of neuroprotection, and of cell therapeutic approaches in C 2009 by John animal models of PD. Curr. Protoc. Stem Cell Biol. 11:2D.5.1-2D.5.21.  Wiley & Sons, Inc. Keywords: stem cells r cell and tissue culture r neuroscience r isolation r puriÞcation r separation r cell and developmental biology r cell therapy

INTRODUCTION This unit describes the dissection of the rat or mouse fetal ventral midbrain (VM) region (Basic Protocol 1) and the generation of dopamine (DA) neuronal cell cultures (Basic Protocols 2 and 3 and Alternate Protocols 1 and 2). Additionally, there is a description of analytical readouts (Support Protocol). These procedures have been applied previously in numerous in vitro and in vivo paradigms (Bjorklund et al., 1983; Nikkhah et al., 1994; Haque et al., 1997; Timmer et al., 2006). The detailed description and the synopsis of updated experimental concepts in this Þeld provided here may help promote a broader application and facilitate the study of midbrain DA neurons in the context of neural development and therapeutic models.

STRATEGIC PLANNING A ßow diagram of experimental options for study design is provided in Figure 2D.5.1. Due to the availability of numerous midbrain-relevant transgenic mice, mouse primary VM tissue can be considered advantageous for many experimental paradigms. This facilitates numerous functional neurobiological in vitro studies, and also comparative studies with murine pluripotent cell sources (Yurek and Fletcher-Turner, 2004; Chung et al., 2006; Lin and Isacson, 2006). Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.5.1-2D.5.21 Published online December 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d05s11 C 2009 John Wiley & Sons, Inc. Copyright 

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preparation of cell suspension

VM dissection

organotypic culture

aggregate culture

adherent culture

experimental read-out

Figure 2D.5.1 Overview: Isolation and culture of VM precursors and DA neurons. Dissection of rodent midbrain (see Basic Protocol 1) enables analysis of intact VM neural tissue for organotypic culture (see Basic Protocol 3), as well as gentle dissociation into single-cell suspensions (see Basic Protocol 2). Cell culture options include expansion and/or differentiation as three-dimensionalaggregate cultures (see Alternate Protocol 1), or as adherent monolayer cultures (see Alternate Protocol 2). Subsequent detailed analysis of DA neuronal phenotype is customized for the specific experimental paradigm at hand (see Support Protocol).

For transplantation studies, rat tissue has been more frequently applied, given the wide array of standardized behavioral tests in rat Parkinson’s Disease (PD) models (Dunnett, 1994). For immediate differentiation, resulting in maturing DA cultures within 1 to 3 days, use tissue from embryonic day 14 (E14) rats or E13 mice (DA neuronal culture). When a short-term expansion step is included (VM precursor culture), use tissue from E11.5 to E12 rats or E11 mice according to the subsequent protocols (see Alternate Protocol 2). Depending on the experimental question at hand, VM tissue of either stage can be used for Basic Protocols 2 or 3. NOTE: Experiments involving live animals must conform to national and institutional regulations and must be approved by the Institutional Animal Care and Use Committee (IACUC) or equivalent. Consider the scientiÞc and biomedical rationale for conducting the particular experiment. NOTE: Sterilize the instruments by autoclaving, and place in the aseptically prepared hood/clean dissection area. Ensure that assigned safety containers are available for scalpel blades and for the glass pipet waste. BASIC PROTOCOL 1

DISSECTION OF VENTRAL MESENCEPHALON This protocol is used to isolate the ventral mesencephalon (midbrain) from rodent embryos, embryonic day (E) 11 to E14, using microdissection techniques. Steps are described on how to identify anatomical landmarks, how to remove overlying tissue, and how to dissect the ventral midbrain portion itself. An accompanying video Þle (see Video 1 at http://www.currentprotocols.com/protocol/sc02d05) demonstrates the entire procedure.

Materials Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

C57B6/J mice (The Jackson Laboratory), embryonic day (E) 11 to E13 or Sprague–Dawley rats (Charles River), E11.5 to E14 Hanks balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170), ice cold Dissection buffer (see recipe)

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Microdissecting instruments (sterilized; Fine Science Tools): Small dissecting scissors Medium dissecting scissors Dumont forceps—straight and angled or curved Curved microdissecting scissors Spatula Moria perforated spoon with holes Laminar ßow hood, sterilized by cleaning with 70% ethanol or UV-exposure for 15 min 60-mm and 100-mm round dishes (petri dishes), Þlled with dissection buffer Dissecting microscope (e.g., Leica MZ6 or Zeiss Stemi 2000) Curved scalpel blade (e.g., BD Bard-Parker no. 23 or 24) 15- and 50-ml conical tubes Collect embryos 1. Using aseptic technique, obtain the uterine horns from a time-pregnant mouse or rat [embryonic age E11.5-12 (rat) or E11.5 (mouse) for VM precursor cells; E14 (rat) or E13 (mouse) for direct DA neuron culture; see Strategic Planning and Fig. 2D.5.1]. 2. Submerge uterine horns in a 100-mm petri dish containing ice-cold, sterile CMFHBSS, and carefully rinse 2 to 3 times with 15 ml ice-cold, sterile CMF-HBSS. NOTE: From this point on work under sterile conditions in a laminar ßow hood, or add antibiotics (penicillin/streptomycin at standard concentrations) to reagents.

3. Transfer to a clean 100-mm petri dish containing dissection buffer. 4. Under a dissection microscope placed in a laminar ßow hood, perform the following steps (steps 5 to 19). Perform the steps in a timely manner, and keep the tissue cooled on ice and immersed in ice-cold buffers throughout the procedure. Follow the dissection sequence as depicted in Figure 2D.5.2. 5. Dissect each embryo from the uterine sac (Fig. 2D.5.2A-F), and remove the amniotic membranes (Fig. 2D.5.2G,H). 6. Using a Morian-type perforated spoon, transfer the embryo to a clean sterile petri dish containing ice-cold dissection buffer. 7. To conÞrm and monitor gestational age, measure and record crown rump length (CRL) of the embryos used. Expect CRL = 5 to 6 mm for early stage embryos (rat E11.5-E12; mouse E11; VM precursor culture). Expect CRL = 10 to 12 mm for later stage embryos (rat E14; mouse E13; DA neuronal culture).

8. Exclude any malformed or otherwise damaged embryos.

Dissect brains 9. Decapitate each fetus using microdissection scissors or a scalpel 10. Identify the central nervous system, and the midbrain region of interest (Fig. 2D.5.2J-M). CAUTION: The tip of the scissors should point away from the brain to avoid damaging the brain. See supplemental video material for a detailed demonstration of the procedure (see Video 1 at http://www.currentprotocols.com/protocol.sc02d05). Somatic Stem Cells

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A

B

C

D

E

F

G

H

I

J

K

L

M

N

O

Figure 2D.5.2 Dissection of the VM region from embryonic rodent brain. (A-F) Isolate the embryo from the uterine sac. (G-H) Free the embryo from any remaining placental and amniotic membranes. (I-L) Decapitate the embryo, and identify anatomical landmarks of the cranial central nervous system. Arrowheads indicate rostral and caudal borders of the midbrain region. Dotted lines outline the contour of forebrain CNS tissue. Arrow in (K) indicates VM region (lateral view). (L-M) Remove the overlying scalp tissue, to isolate the brain (superior view). Cut away the rostral forebrain and the caudal hindbrain regions (dashed lines; lateral view). (N) Open the resulting tube-like structure along the posterior midline (dashed line; coronal view). Arrow indicates anterior midline and VM region. (O) Trim the resulting butterfly-shaped structure, removing ∼2/3 of the posterior/lateral tissue on each side (dashed line; view from ventral midline, tissue flattened). Arrow indicates anterior midline of VM region. Also, see the supplemental Video 1 at http://www.currentprotocols.com/protocol/sc02d05. Abbreviations: fb, forebrain; hb, hindbrain; VM, ventral midbrain.

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11. Carefully isolate the brain (Fig. 2D.5.2N). Holding the tissue with forceps near the forebrain or hindbrain regions (to be discarded in this VM DA cell isolation protocol), dissect away and remove the overlying scalp tissue. IMPORTANT NOTE: Avoid touching and damaging the midbrain region itself throughout the dissection procedure. The forebrain tissue can be processed in an identical manner, e.g., for comparative analysis or alternative studies.

12. Place the isolated brain in a clean 60-mm petri dish containing dissection buffer on ice. 13. Stabilizing the brain with forceps near the forebrain or hindbrain regions, carefully remove the fore- and hindbrain regions using a scalpel or microscissors. Place the rostral cut close to the forebrain vesicles and thalamic region and the caudal cut at the isthmus region (Fig. 2D.5.2M). If working with a scalpel, use the blade as a shield protecting the midbrain region, while removing the unwanted tissue with forceps (see Video 1 at http://www.currentprotocols.com/ protocol/sc02d05).

Dissect the ventral midbrain 14. Using forceps, steady the obtained midbrain tube (Fig. 2D.5.2O), exclusively touching the posterior midbrain region marked by the convex curvature at the dorsal midline. 15. Use small microscissors or the very tip of a curved scalpel blade to gradually dissect open this tube along the dorsal midline. 16. Carefully open the (Fig. 2D.5.2N,O).

now

characteristically

butterßy-shaped

tissue

ßap

17. Use forceps to thoroughly remove any remaining overlying meningeal tissue. Any remaining meningeal tissue can be recognized on the ventral exterior surface by its dense vascularization. When using the VM tissue for transplantation studies, such contaminating cells can promote unwanted immune reactions (Chen and Palmer, 2008). In vitro, the Þbroblast and endothelial cell types can overgrow and decrease the purity of the primary DA cultures.

18. Trim the outermost areas, i.e., the most dorsal parts of the midbrain tube, by dissecting away approximately two thirds of the tissue on each side (Fig. 2D.5.2O; i.e., approximately lateral/posterior to the sulcus limitans as an anatomical landmark). 19. Transfer the resulting tissue piece with dimensions of ∼0.3 × 1.0–mm into a conical tube containing cold dissection buffer kept on ice. Use ∼0.2 to 0.5 ml buffer volume per each VM. Note that, while immediate use is highly recommended, VM DA cell tissue pieces can in principle be stored up to 2 days at 4◦ C, in a hibernation medium and still yield viable VM DA cultures (Nikkhah et al., 1995). This strategy is utilized in clinical cell therapeutic paradigms, where human fetal VM DA tissue has been kept in hibernation medium supplemented with glial cell-derived neurotrophic factor (GDNF) prior to transplantation into patients (Mendez et al., 2005, 2008).

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BASIC PROTOCOL 2

PREPARATION OF CELL SUSPENSION This protocol is used for generation of a cell suspension from neural tissue (dissected in Basic Protocol 1). Such primary midbrain-derived cell preparations are used for in vitro culture or directly for transplantation assays. NOTE: For VM organ culture, skip this protocol and proceed directly as described in Basic Protocol 3. To obtain a cell suspension for use in three-dimensional aggregate cultures (Alternate Protocol 1) and/or adherent in vitro culture systems (Alternate Protocol 2), proceed as follows.

Materials Ventral midbrain tissue (Basic Protocol 1) Dissection medium (see recipe) Dissociation medium (see recipe) or trypsin 0.05% (w/v)/ EDTA (Invitrogen, cat. no. 25300) containing 0.2% (w/v) DNase I (see recipe) or Accutase (Innovative Cell Technologies, cat. no. AT104) or TrypLE Express (Invitrogen, cat. no. 12605) Heat-inactivated fetal bovine serum (FBS; Hyclone, cat. no. SH30070) Expansion medium (see recipe) Differentiation medium (see recipe) Trypan blue (Invitrogen, cat. no. 15250) or acridine orange/ethidium bromide solution (see recipe) 15-ml conical tubes Laminar ßow hood 37◦ C water bath Sterile Þre-polished 9-in. Pasteur pipets (see recipe) 200- and 1000-μl plastic tips and pipettors 70-μm cell strainer (BD, cat. no. 352350) or round bottom tube with 35-μm cell strainer caps (BD, cat. no. 352235) 1.5-ml microcentrifuge tubes Benchtop centrifuge Hemacytometer Microscope for viability dye detection (trypan blue: light microscope with bright Þeld or phase contrast; acridine orange/ethidium bromide: ßuorescence microscope with UV excitation and Þlters appropriate for simultaneous red-green channel detection; emission max for DNA is 526 nm, for RNA 650 nm) Additional reagents and equipment for determining the cell concentration and viability using trypan blue (UNIT 1C.3) Dissociate cells 1. Wash the obtained tissue pieces (Basic Protocol 1) in cold dissection buffer (e.g., 15 ml buffer in a 15-ml conical tube), by letting the tissue pieces sink down in the conical tube, pipetting off the medium, and Þlling the tube with fresh buffer. 2. Pipet off the buffer, and add 1 ml (per 10 midbrain tissue pieces) of the solution to be used for dissociation. Use either dissociation medium, or alternatively 0.05% trypsin/EDTA with 0.2% (w/v) DNase added, Accutase, or TrypLE Express. Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

To reduce the clumping of cell suspensions due to sticky nucleic acids released by damaged cells during dissociation, the addition of DNase during mechanical dissection and when using trypsin/EDTA or TrypLE Express is highly recommended (Panchision et al., 2007; Pruszak et al., 2007). The commercially available preparation Accutase already contains DNase activity.

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3. For enzymatic digestion, incubate 3 to 15 min at 37◦ C. Use visual control and determine the optimal duration by test trituration. Avoid over-digestion, and inactivate with 10% fetal bovine serum if required (for trypsin/EDTA digestion). This is a critical step. Obtaining a viable neural cell suspension is a prerequisite for the subsequent culture protocols.

4. Using Þre-polished Pasteur pipets with decreasing diameter, gently dissociate the tissue pieces, ∼20 times total. Alternatively, trituration may be performed using Þrst a 1000-μl pipettor, followed by trituration with a 200-μl pipettor. Avoid excessive formation of air bubbles during mechanical dissociation of VM tissue, as those will reduce cell viability.

5. In case major tissue chunks remain in the solution, selectively triturate those pieces separately. Consider discarding some tissue, rather than compromising the major part of the cell suspension due to mechanical dissociation. For an optional Þltering step, pipet the obtained cell suspension through a cell strainer cap or through a 35- to 70-μm mesh. To minimize loss of cells due to this step, subsequently ßush the Þlter membrane with a small volume of medium.

6. Centrifuge 3 to 5 min at 200 × g, 4◦ C . Pipet off supernatant. 7. Resuspend in plating medium (either expansion medium or differentiation medium). The amount of plating medium varies (e.g., 200 μl per 10 midbrain pieces originally isolated).

Determine cell number and viability 8. Determine the cell concentration and the viability of an aliquot of the cell suspension, using a classic dye exclusion method (trypan blue; UNIT 1C.3) or DNA/RNA labeling techniques (acridine orange/ethidium bromide). Use a small sample of the obtained cell suspension and dilute it with the viability test dye, at a ratio of 1:10 (e.g., 1 or 5 μl cell suspension and 9 or 45 μl of dye, respectively). 9. After gentle mixing, transfer the sample solution with a fresh pipet tip to a hemacytometer chamber for visual inspection and quantiÞcation under a microscope. 10. Determine the viability and calculate the cell concentration. In trypan blue staining, dead cells will take up the dye and will appear blue under a light microscope. With acridine orange/ethidium bromide, live cells appear green (acridine orange due to labeling of RNA), while dead cells are labeled red (due to acridine orange and ethidium bromide labeling of DNA) upon UV-excitation and detection in the 520 to 650 nm range on a ßuorescence microscope. Cell viability needs to be higher than 80%, and should routinely range from 95% to 100% (see Fig. 2D.5.4A). Note that DA neurons are among the most fragile cells in the solution, and while cultures will contain neuronal cell types after relatively harsh treatment, the DA numbers will be low. Use the live cells counted for calculating the cell concentration.

11. Keep the cell suspension on ice or at 4◦ C until use. Before proceeding with either the three-dimensional aggregate (Alternate Protocol 1) or the adherent culture steps (Alternate Protocol 2), or with transplantation experiments, consider including puriÞcation methodologies recently optimized for neural cell suspensions such as ßuorescence-activated cell sorting (FACS) or immunomagnetic cell separation (MACS; Kerr et al., 1994; Ono et al., 2007; Panchision et al., 2007; Pruszak et al., 2007); also see the ßow chart in Fig. 2D.5.1).

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BASIC PROTOCOL 3

MIDBRAIN NEURAL CULTURE: ORGANOTYPIC CULTURE This protocol outlines the detailed procedures for in vitro culture of VM precursor and DA neuronal cell types. One option is to culture midbrain tissue obtained in Basic Protocol 1 as intact organotypic cultures on a membranous insert in the well. This preserves, to some extent, a physiological cellular context. An alternative option is the formation of three-dimensional aggregate cultures. Sectioning such spherical aggregates provides a good readout of cells grown in near-physiological conditions (Alternate Protocol 1). The most commonly applied option is the culture of VM neural precursors or DA neurons on permissive substrates such as laminin and/or poly-L-ornithine (Alternate Protocol 2). NOTE: Either early stage (VM precursors, E11.5-12 rats, E11 mice) or later stage fetal tissue (DA neurons, E14 rats, E13 mice ) can be used. However, the efÞciency for a short-term expansion step is limited to the early stage VM precursors (E11.5-12 rat; E11 mouse; CRL ≈5 to 6 mm. See Alternate Protocol 2).

Materials Ventral midbrain tissue pieces (Basic Protocol 1) Differentiation medium (see recipe) 4% (w/v) paraformaldehyde (PFA) solution Laminar ßow hood Pasteur pipet with a Þre-polished widened oriÞce (see recipe) or curved forceps Forceps or tungsten needles Millicell cell culture inserts (for six-wells; e.g., Millipore, cat. no. PICM0RG50) 6- and 24-well tissue culture plates (e.g., Fisher, Falcon or Nunc) 37◦ C water bath Scalpel Perform organotypic cultures 1. After isolating the midbrain tissue (characteristic “butterßy” structure, Basic Protocol 1; see Video 1 at http://www.currentprotocols.com/protocol/sc02d05; see Fig. 2D.5.2O), carefully transfer the intact VM tissue piece to a membrane-covered tissue culture insert, either by using a Pasteur pipet with a Þre-polished widened oriÞce, or by carrying the tissue in a liquid droplet between the tips of the branches of a curved pair of forceps (Fig. 2D.5.3A). 2. Using forceps or tungsten needles, gently ßatten the tissue and orient it towards the central area of the insert, away from the edges. 3. Using a pipet, very carefully remove any remaining ßuid on the top surface of the insert’s membrane, to ensure adherence to the membrane and to avoid ßoating of the tissue. This is a critical step to avoid ßoating of the tissue.

4. Place the insert into the well of a six-well culture plate Þlled with 1.5 ml differentiation medium. Ensure that no air bubbles remain below the insert’s membrane. 5. Culture for up to 10 days, while changing medium every two days in the compartment below the insert.

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6. For staining and analysis of this in vitro system, Þx the tissue in the well 20 min at room temperature with 4% paraformaldehyde solution, and carefully cut out the membrane from the insert using a scalpel blade. 7. Transfer the membrane piece with the VM tissue attached to a smaller well format (4-well or 24-well) for immunocytochemistry. 8. For further speciÞcs regarding staining and analysis, refer to Support Protocol. Current Protocols in Stem Cell Biology

A

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Figure 2D.5.3 VM region organotypic culture. (A) Transfer of intact VM tissue onto tissue culture insert. (B) Precursor stage (rat E12) VM region organ culture stained for the DA marker TH. (C) Rat E12 VM region organotypic culture after 2 days in vitro, stained for TH. (D) Higher magnification of VM DA neurons in intact VM region tissue culture. Abbreviations: TH, tyrosine hydroxylase; DA, dopaminergic; VM, ventral midbrain; E12, embryonic day 12; div, days in vitro. Scale bars = 100 μm.

MIDBRAIN NEURAL CULTURE: THREE-DIMENSIONAL AGGREGATE CULTURE

ALTERNATE PROTOCOL 1

An alternative for culturing midbrain neural cells after dissociation is the three dimensional neural culture (see Fig. 2D.5.4).

Additional Materials (also see Basic Protocols 2 and 3) VM cell suspension (Basic Protocol 2) Differentiation or expansion medium (see recipes) 4% (w/v) paraformaldehyde solution 15% (w/v) agar gel 15-ml conical tubes Shaker/roller tube system (e.g., Miltenyi Biotec, cat. no. 130-090-753, MACSmix Tube Rotator) HumidiÞed tissue culture incubator (37◦ C, 5% CO2 ), preferably including low O2 option Vibratome 1. Dilute the obtained cell suspension appropriately. For aggregate culture, transfer ∼1–5 × 105 live VM cells dissolved in 5 ml medium into a 15-ml conical tube. Somatic Stem Cells

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A

AO/EthBr

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Figure 2D.5.4 Three-dimensional-aggregate culture. (A) Viability of VM cell suspensions for three-dimensional-aggregate cultures, adherent culture systems, and transplantation studies alike is determined by viability dyes. Here, acridine orange/ethidium bromide (fluorescent image showing live cells in green; upper panel). Lower panel: phase contrast image of the identical field. (B) Aggregate formed after 7 days in vitro in the roller tube system. (C) Aggregate cultures stained for nuclear marker DAPI (tightly packed), TuJ1 neuronal marker, dense fiber network surrounding. (D) Aggregate cultures stained for tyrosine hydroxylase. Inset: higher magnification. Sections of aggregates cut on a vibratome after embedding in agarose are shown; C and D display identical areas. Abbreviations: AO, acridine orange; EthBr, ethidium bromide; TuJ1, neuronal marker beta-III tubulin; DAPI, nuclear marker; TH, tyrosine hydroxylase.

2. Place the tube into a shaker/roller tube system that enables a steady rocking or turning motion of about 30 cycles per min, inside a tissue culture incubator. NOTE: Set up the roller tube system (Alternate Protocol 1) inside the tissue culture incubator. Arrange in such a way that liquid in the tubes will not leak out (if needed, position the system in a tilted but stable manner).

3. Culture for 3 to 10 days. Change medium every other day, by letting the cells settle/sink to the bottom, placing the tube vertically for ∼10 min, and then carefully removing the supernatant, leaving a small volume of medium to avoid cell loss. After 3 to 5 days, macroscopically visible clusters of cells will have formed.

4. For staining and analysis of this in vitro system, Þx the cells in the tube with 4% paraformaldehyde solution. Embed each aggregate in a 15% agar gel. Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

5. Carefully cut each aggregate using a vibratome into sections of ∼20-μm thickness. Those sections can be further processed according to standard procedures such as immunocytochemistry. For further speciÞcs regarding staining and analysis, refer to Support Protocol.

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MIDBRAIN NEURAL CULTURE: ADHESION CULTURE Adhesion cultures of VM cells enable the three-dimensional culture of dissociated cell suspensions (see Fig. 2D.5.5).

ALTERNATE PROTOCOL 2

Materials VM cell suspension (Basic Protocol 2) Expansion medium (see recipe) Differentiation medium (see recipe) 24-well tissue culture plates Laminin/poly-L-ornithine coated 12-mm coverslips (see recipe) 100- or 200-μl pipets HumidiÞed tissue culture incubator (37◦ C, 5% CO2 ), preferably including low O2 option Dilute the obtained cell suspension 1a. For expansion of VM precursor cells (E11.5-E12 rats; E11 mice): Plate the cells in 24-well tissue culture plates at densities of 1–5 × 104 cells per cm2 in 0.5 to 1.5 ml expansion medium including 20 ng/ml bFGF as a mitogen. 1b. For differentiation of DA neuronal cell suspension (E14 rats; E13 mice): Plate at densities of 2–5 × 105 cells per cm2 in 0.5 to 1.5 ml differentiation medium. 2. To maximize the yield, plate cells as 30-μl droplet cultures, using a 100-μl or 200-μl pipet, on a freshly coated, washed, and brießy air-dried coverslips. A 30-μl drop covers an area of ∼1 cm2 .

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Figure 2D.5.5 Adherent culture system. (A) VM DA precursors (E11-12) at 1 day in vitro after plating, and (B) at 7 days in vitro of expansion with bFGF. (C) VM DA precursors during the expansion phase stain positive for Nestin (red), a minor fraction of cells stains positive for beta-III-tubulin (green). Blue = nuclear Hoechst stain. (D) Expansion and proliferative capacity is monitored by BrdU incorporation assays: here ranging from 44.2% BrdU+ cells at 1 day in vitro, to over 46.4% at 5 days in vitro to 30.8% at 7 days in vitro. Error bars indicate SEM; three independent experiments. (E) Differentiation of DA neurons is induced subsequent to in vitro expansion or alternatively immediately after VM dissection from older embryos (E14), forming a dense network of neuronal processes, staining positive for neuronal markers such as beta-III-tubulin (TuJ1, green; F, G) and dopaminergic markers such as tyrosine hydroxylase (TH, red; H).

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Calculate the cell concentration of the cell suspension accordingly. For example, for plating of 20 coverslips of E11.5 VM precursor cells as 30-μl droplets (area ≈1-cm2 ), at a density of 20,000 cells per cm2 , 400,000 cells suspended in 600 μl expansion medium are needed. Consider loss due to pipetting and measuring errors, and prepare cell suspension in surplus.

3. After gentle mixing of the cell suspension (avoid additional trituration), position a 30-μl cell suspension droplet onto the center of a coverslip in a 24-well plate. 4. Repeat until all wells are plated with the available cell suspension in the same manner. 5. Using caution not to spill the droplets, place the tissue culture plate into an incubator for ∼0.5 hr. 6. Carefully Þll up the wells to a volume of 0.5 to 1.3 ml with expansion or differentiation medium per 24-well. When too much debris is present, a gentle washing step, adding fresh medium, can be performed at this stage. For VM precursor cultures use expansion medium, for DA neuronal cultures use differentiation medium. Ensure that the coverslips are entirely covered by medium. Remove potential air bubbles under the coverslips, as those may later cause ßoating of the coverslips, and drying of the surface area.

7. For the VM expansion cultures, change medium from expansion medium to differentiation medium after 2 to 4 days. 8. Culture for 3 to 10 days, changing the medium every other day. Depending on the cell density and resulting metabolic turnover, daily medium changes should be considered at prolonged stages in culture (monitor pH/phenol red indicator property of medium). DA neuronal primary cultures are very sensitive. Perform the medium changes rapidly and carefully, to avoid letting any well dry.

9. Process for analysis (see Support Protocol). SUPPORT PROTOCOL

ANALYSIS OF VM NEURAL PRECURSORS AND DA NEURONS This section summarizes and discusses the analytical readout options available to study VM neural precursors and DA neurons (see Fig. 2D.5.6).

Materials

Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

Dulbecco’s phosphate-buffered saline (DPBS) Mg+, Ca+-free (CMF-DPBS; Invitrogen, cat. no. 14190) Antibodies typically used in a basic VM DA differentiation: Sheep anti-TH (1:1,000; Pel-Freez) Mouse anti-nestin (1:100; Millipore/Chemicon) Rabbit anti-TuJ1 (Covance 1:1000) Mouse anti-MAP2 (Millipore/Chemicon 1:500) Mouse anti-Pitx3 (Zymed 1:1000) Rabbit anti-Pitx3 (1:250; Invitrogen) Rabbit anti-glial Þbrillary acidic protein (1:500; Dako) Rabbit anti-Nurr1 (E-20; 1:300; Santa Cruz Biotechnology) Mouse anti-engrailed 1 (clone 4G11; 1:40) Rabbit anti-ki67 (1:2,000; Novocastra/Vector Laboratories) Rabbit anti-DAT (1:1000; Millipore/Chemicon) Corresponding secondary antibodies Pipets

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Microscope for cell analysis Vibratome for sectioning of three-dimensional-aggregate cultures (Leica VT1000 S; Basic Protocol 3) Analysis of midbrain neural precursor and DA neuronal cultures Staining protocols include immunohistochemical, e.g., immunoperoxidase-based, and immunoßuorescence techniques. For a detailed description of these procedures, refer for example, to Glynn and McAllister, 2006; Kim et al., 2006; or Hoffman et al., 2008. Commonly used antibodies for a basic analysis of neural differentiation include those against the neural precursor marker nestin, and neuronal markers such as TuJ1 (beta-3tubulin), MAP2, or Tau. Proliferation assays may be performed using bromodeoxyuridine (BrdU) incorporation to determine the expansion potential of neural precursors. Astroglial differentiation can be documented by staining for glial Þbrillary acidic protein (GFAP). Basic analysis of midbrain DA phenotype includes staining for the catecholaminergic marker tyrosine hydroxylase (TH), co-labeled with DA transcription factors such as Pitx3 and/ or Nurr1. The A9 DA neurons speciÞcally express the marker GIRK2. Stained neurons in vitro may be analyzed with respect to neurite outgrowth, directed targeting, or the inßuence of co-culture conditions, e.g., with mesencephalic glial cell types. Supplementing such immunoßuorescence and morphological studies, phenotypic characterization is also done by gene expression analysis via RT-PCR, and by protein analysis (Western blot; Sonntag et al., 2007). SpeciÞc subpopulations can be analyzed by using techniques such as laser capture microdissection (Espina et al., 2006), or ßow cytometric analysis of cell subsets labeled with antibodies or transgenic ßuorescent markers (Pruszak et al., 2007).

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TuJ1/Hoechst

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Figure 2D.5.6 Options for analytical readout of VM DA neurons. Immunocytochemical assays include measures of neurite outgrowth and targeting studies (A), and/ or co-culture assays of VM DA neurons, here using astroglial feeder cells (B). Detailed analysis of specific DA neuronal subsets is achieved by isolating fixed DA neurons using laser capture microdissection (LCM) (C). Fluorescence-activated cell sorting (FACS) methods optimized for fragile neural cell types enables isolation of viable VM DA neurons for further in vitro and in vivo analysis in pharmacological, toxicological, and cell transplantation assays (D,E).

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Functional aspects of live VM DA neurons in vitro may be studied through electrophysiological analysis (Schlesinger et al., 2004). Furthermore, HPLC serves to detect dopamine release and dopamine metabolites such as DOPAC in media supernatants and/or cell samples as additional parameters for evaluating the extent of DA differentiation and the functionality of the cultured cells. Typically, the latter test is performed under basal conditions and then after stimulation, e.g., with 56 mM potassium chloride solution. See Studer et al. (1998) and Chung et al. (2002) for details. Finally, in vivo transplantation of VM DA neurons, commonly ectopically into the striatum of rodent animal models, enables the study of integration of VM DA neurons into the host circuitry by histological, as well as behavioral analyses (Vinuela et al., 2008; Nikkhah et al., 2009).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Acridine orange/ethidium bromide solution Prepare a solution of 5 μg/ml acridine orange (Sigma, cat. no. A8097) and 5 μg/ml ethidium bromide (Sigma, cat. no. E151) in phosphate-buffered saline (e.g., CMF-DPBS; Invitrogen, cat. no. 14190). Store up to 12 months at 4◦ C. CAUTION: Acridine orange/ethidium bromide solution is mutagenic and is a health hazard. This solution (10-fold) can be prepared and stored at 4◦ C, protected from light, for 3 months).

Ascorbic acid (AA) stock solution (1000×) Dissolve ascorbic acid (Sigma, cat. no. A4034) in phosphate-buffered saline (CMFDPBS; Invitrogen, cat. no. 14190) to prepare a 200 mM stock solution, and Þltersterilize using a 0.22-μm Þlter. Protect from light. Store up to 12 months at −20◦ C. Basic Þbroblast growth factor (bFGF) stock solution Dissolve bFGF (Invitrogen, cat. no. PMG0034) in phosphate-buffered saline (CMFDPBS; Invitrogen, cat. no. 14190) at 2 μg/ml. Divide into 50-μl aliquots and store up to 6 months at −20◦ C. Differentiation medium Neurobasal medium (Invitrogen, cat. no. 21103) L-glutamine (Invitrogen, cat. no. 21051-016) 50× B27 (Invitrogen, cat. no. 17504) 1% (v/v) heat-inactivated fetal bovine serum (FBS; Hyclone, cat. no. SH30070) Add 100 μM ascorbic acid (see recipe) immediately before use Store up to 7 days at 4◦ C ModiÞcations known to enhance the DA fraction include the addition of growth factors (see recipe) such as GDNF (Costantini and Isacson, 2000), FGF20 (Ohmachi et al., 2003), or TGF3beta to the differentiation medium.

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Dissection buffer Hanks balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170) 20 mM D-glucose (Sigma, cat. no. G8270) Penicillin/streptomycin (Invitrogen, cat. no. 15140; use standard concentrations, as indicated by the supplier) Just before use, add 100 μM ascorbic acid (see recipe) Shelf life: 1 week at 4◦ C Current Protocols in Stem Cell Biology

Dissociation medium DMEM/F12 (Invitrogen, cat. no. 11320) 0.05% (w/v) DNase (Sigma, cat. no. D5025) 50× B27 (Invitrogen, cat. no. 17504) Prepare fresh DNase I stock Dissolve DNase I (Sigma, cat. no. D5025) at 0.2 mg/ml in Hanks’ balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170). Divide into aliquots and store up to 12 months at −20◦ C. Prepare sufÞcient quantity by dissolving 0.0024 g of DNase in 12 ml of HBSS/glucose (see recipe; this large volume allows for several rinses and trituration). Sterile Þlter using a 0.22-μm Þlter and keep on ice; then use or freeze.

Expansion medium DMEM/F12 (Invitrogen, cat. no. 11320) 100× N2 (Stem Cell Technologies, cat. no. 07152) Add 20 ng/ml bFGF (see recipe) immediately before use Store up to 2 weeks at 4◦ C Refer to Bouvier and Mytilineou (1995) and Studer et al. (1998) (the original papers introducing bFGF supplementation in the expansion medium).

Fire-polished glass pipets Prepare a set of Þve to ten Þre-polished pipets over a small ßame (alcohol or natural gas burner), such that the edges are smoothed and with decreasing aperture diameter starting at ∼1 mm (Schnitzler et al., 2008). Autoclave.

Growth factors Prepare as 1000× stocks and divide into aliquots. Store up to 6 months at −80◦ C. Add to the expansion or differentiation medium (see recipes) just before plating or medium change.

HBSS/glucose solution Dissolve 1.08 g D-glucose (Sigma, cat. no. G8270) in 500 ml Hanks’ balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170) to prepare a 20 mM glucose solution and Þlter-sterilize using a 0.22-μm Þlter. Store up to 2 weeks at 4◦ C.

Laminin solution Thaw laminin stock (Becton-Dickinson, cat. no. 354232) slowly to avoid gelatinization. Prepare on ice, using cooled pipets (kept at −20◦ C) to a Þnal concentration of 1 μg/ml in CMF-DPBS. Use immediately. CAUTION: Laminin rapidly adheres to surfaces (i.e., vessels, pipets, tips) if not kept ice-cold throughout preparation.

Poly-L-ornithine 0.01% (w/v) solution (Sigma, cat. no. P4957) Þnal. Dilute stock solution of polyL-ornithine [5 mg/ml in Dulbecco’s phosphate-buffered saline (DPBS) Mg++ , Ca++ -free (CMF-DPBS; Invitrogen, cat. no. 14190] and Þlter-sterilize using a 0.22-μm Þlter. Store up to 2 weeks at 4◦ C.

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Poly-L-ornithine/laminin-coated plates Coat plates (six-wells or 24-wells with glass coverslips) sequentially with poly-Lornithine (15 μg/ml; see recipe) and then laminin (1 μg/ml; see recipe) solutions. Throughout, ensure to add sufÞcient quantity of coating solutions or washing buffer to cover the surface of culture plates. First, incubate with poly-L-ornithine solution (15 μg/ml; see recipe) for 2 hr at 37◦ C or overnight at room temperature. Remove solution, wash with deionized water. Second, add freshly prepared ice-cold laminin solution (1 μg/ml; see recipe) using cooled pipets, and incubate for 2 hr at 37◦ C or overnight at room temperature. Remove solution, wash three times with 2 ml deionized water. In our hands, we obtain most consistent results using the coated plates right away, either “wet” or after brief (15 min) air drying in the hood. Note that dry coverslips are required for droplet culture (see Alternate Protocol 2).

COMMENTARY Background Information

Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

Dopamine (DA) neurons of the substantia nigra region (pars compacta), A9 region, in the ventral midbrain (VM) play a critical role in the initiation of movement, as well as in behavior, motivation, emotion, and cognition through their contribution to basal ganglia neural circuitry (Ungerstedt, 1976; Fibiger and Phillips, 1988; Koob and Swerdlow, 1988; Perrone-Capano and di Porzio, 1996). Their progressive degeneration is a major feature of Parkinson’s disease (Olanow, 2007), leading to signiÞcant motor disability (Weintraub et al., 2008). DA neurons are generated from midbrain (mesencephalic) precursor cells, which originate from the neuroepithelial layer in the ventral neural tube of that region (Prakash and Wurst, 2006; Smits et al., 2006; Ono et al., 2007; Smidt and Burbach, 2007). Transcription factors relevant for the in vivo speciÞcation of DA neurons have been identiÞed in signiÞcant detail utilizing wild-type and DA-relevant transgenic mouse strains, such as Pitx3 (Smidt et al., 2004a,b), Lmx1b (Asbreuk et al., 2002), Otx2 (Puelles et al., 2004; Borgkvist et al., 2006), Nurr1 (Saucedo-Cardenas et al., 1998; Smits et al., 2003), Fox2a (Ferri et al., 2007), and Ngn2 (Andersson et al., 2006; Thompson et al., 2006). Functional DA neurons have also been generated from embryonic stem cells (Lee et al., 2000; Barberi et al., 2003; Perrier et al., 2004; Chung et al., 2006), and more recently mouse induced pluripotent stem (iPS) cells (Wernig et al., 2008). In the induction of DA neurons from such pluripotent cell sources, a major fraction has been shown to express DA-speciÞc markers, and the proofof-principle of restoring function in PD animal models has been demonstrated (Bjorklund

et al., 2002; Sonntag et al., 2007; Wernig et al., 2008). However, such DA patterning efforts from pluripotent sources require further optimization and Þne-tuning (Pruszak and Isacson, 2009), as so far only a smaller fraction of DA neurons derived from pluripotent cell sources may actually be patterned toward a true equivalent of the physiological phenotype (Hedlund et al., 2008). Thus, DA cells obtained from primary VM tissue represent a physiologically derived gold standard for studies of DA neuronal features, such as lineage development (Thompson et al., 2006; Ono et al., 2007), selective vulnerability (Chung et al., 2005, 2007), and functionality in behavioral (Dunnett, 1994; Klein et al., 2007) and electrophysiological assays (Geracitano et al., 2005; Rick et al., 2006). They provide a means for pharmacological and toxicity testing (SalthunLassalle et al., 2004; Chung et al., 2005), and serve as a valuable reference point for the derivation of DA neurons from pluripotent stem cells (Lin and Isacson, 2006).

Critical Parameters and Troubleshooting Culturing DA neurons Low yield of DA neurons is often a result of (1) too generous dissection of VM region, (2) too harsh trituration of VM tissue or (3) too long an exposure to enzymatic digestion during the cell preparation process. ModiÞcations known to enhance the DA fraction include the addition of GDNF (Costantini and Isacson, 2000), FGF20 (Ohmachi et al., 2003), or TGF3beta to the differentiation medium. Viability Similarly, low overall viability is usually due to harsh trituration generating many air

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bubbles, or extended enzymatic digestion during the cell preparation process. Use a Þrepolished Pasteur pipet with gentle trituration. Adherence Poor adherence after plating suggests that the quality of the coating is not sufÞcient to induce or support DA neuronal growth. Prepare coating solutions freshly, keep them cool, and apply them promptly. If necessary, precoating of poly-L-ornithine/laminin-coated dishes with 10% FBS in PBS for ∼10 min prior to plating can improve adherence (remove FBS solution from the well before plating cells). Gently wash and change to fresh medium the following day, particularly for the VM expansion protocol, as serum components will induce differentiation. Plating density Higher-density plating (e.g., 0.5 × 106 cells/cm2 ) will often yield better quality cultures with improved appearance; however, analysis on a single-cell level can be difÞcult in such dense cultures. Note that more frequent medium changes and increasing the medium volume per well will be required for culture plated at higher densities. Contamination Under aseptic conditions, VM DA cultures can be grown without antibiotics, which is the preferred method. Antibiotics can foster bacterial resistance and could interfere with physiological tests performed. Nevertheless, penicillin-streptomycin is still widely applied in tissue culture as a preventative measure. Other suspected contamination of VM DA cultures can be successfully treated with additional supplementation of Normocin (Invivogen, cat. no. ant-nr-o) to the medium. Mycoplasma spp. are generally less of an issue in primary culture, but cross-contamination from existing cell-line work in the laboratory cannot be excluded. After positive testing of cell cultures for Mycoplasma spp., eradication can be attempted using enroßoxacin (Sigma, cat. no. 17849) at a Þnal concentration of 25 μg/ml.

Anticipated Results Using this detailed guideline and demonstration, the procedures can be easily followed by a researcher without prior VM DA isolation experience. In roller tube aggregation cultures (Alternate Protocol 1), macroscopically visible aggregates should form within two divisions. Those can be monitored under a

standard microscope at 4× to 10× magniÞcation by focusing on the aggregate that has sunk to the bottom of the conical tube. Using the adherent culture system (Alternate Protocol 2) for expansion of VM precursor cells (rat E11 embryos; CRL ∼6 mm), a net expansion of 3- to 5-fold over the course of 3 days can be expected. bFGF is used as an efÞcient mitogen for VM DA precursors (Bouvier and Mytilineou, 1995; Studer et al., 1998). On average, 60% of cells at this stage are Nestin+ when cultured with bFGF-containing medium as described here (ranging from up to 85% of cells at day 2 in culture to ∼40% after 7 days of expansion). Approximately 45% of cells incorporate the proliferation marker bromodeoxyuridine (BrdU) during the expansion phase, when incubated under the conditions described above. For VM expansion cultures, E11 embryos (CRL ∼6 mm) yield the highest efÞciency/number of cells. Note that longer expansion times result in a decrease in proliferative capacity (e.g., 30% BrdU+ cells at 7 div). Expect ∼90% of cells to be positive for markers of neuronal phenotypes at this stage. Contaminating astrocytes, positive for glial markers such as GFAP, are routinely <5% in such short-term cultures. To induce differentiation, bFGF is omitted from the culture medium. After expansion and subsequent differentiation as described above, the cultures are composed of mainly neuronal cells (TuJ1+ , MAP2+ ), ∼30% of which stain positive for tyrosine hydroxylase (when grown under low oxygen conditions —3% to 5% oxygen). Expect numbers in the range of 10% to 15% of TH over TuJ1 when using standard CO2 incubators without this low oxygen option. For VM DA differentiation stage, after 7 days in vitro, cells are primarily TuJ1+ and MAP2+ . The initial VM precursor expansion period can be extended, but it is known that DA differentiation capacity decreases, and extended culture in vitro may enhance tissue culture artifacts and result in less physiological cells over time. Culture conditions can be further modiÞed by addition of factors beneÞcial for DA differentiation and/ or survival, such as FGF-20 (Ohmachi et al., 2003; Grothe et al., 2004), TGF3b (Roussa et al., 2006). Additional approaches have been developed to optimize the purity of neural cell suspension cultures (Pruszak et al., 2007). For VM DA experimental systems, this has been adapted by Ono et al., using FACS for the corin antigen identiÞed on VM DA precursors (Ono

Somatic Stem Cells

2D.5.17 Current Protocols in Stem Cell Biology

Supplement 11

Table 2D.5.1 Time Requirements of Methods Described in this Unit

Procedure

Time to complete

Dissection of ventral mesencephalon

1 to 2.5 hr

Preparation of cell suspension

0.5 hr

Midbrain neural culture

3 to 10 days

VM region organ culture

3 to 10 days

Three-dimensional-aggregate culture

3 to 10 days

Adherent culture

3 to 10 days

Plating of VM neural cell cultures

1 hr

Expansion of VM precursors

3 days

Differentiation of DA neurons

3 to 10 days

Analysis of cultures

2 to 3 days

Immunocytochemistry

2 days

Microscopic analysis

1 day

Transplantation

4-6 hr

In vivo analysis

4-12 weeks

Laser capture microdissection

6-8 hr

Flow cytometric analysis

4-6 hr

et al., 2007). Moreover, combinations of the procedures presented here make the described method highly versatile. For example, consider co-culture systems of three-dimensional aggregates (Alternate Protocol 1) by transferring to membranes with VM region organotypic cultures (Basic Protocol 3) or to adherent culture systems (Alternate Protocol 2) to study cell-cell and tissue interactions.

Time Considerations For time requirements of the various procedures of this unit, see Table 2D.5.1.

Acknowledgements This work was supported by grant DFG (Ni330) from the German Research Council (DFG) to GN. We thank A. Bader for generously sharing reagents and equipment during early stages of this project. We remain grateful to Z. Gong, J. Maciaczyk, C. Mauth, and A. Vinuela for fruitful collegial discussions, as well as methodological support.

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Current Protocols in Stem Cell Biology

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Glynn, M.W. and McAllister, A.K. 2006. Immunocytochemistry and quantiÞcation of protein colocalization in cultured neurons. Nat. Protoc. 1:1287-1296.

Mendez, I., Vi˜nuela, A., Astradsson, A., Mukhida, K., Hallett, P., Robertson, H., Tierney, T., Holness, R., Dagher, A., Trojanowski, J.Q., and Isacson, O. 2008. Dopamine neurons implanted into people with Parkinson’s disease survive without pathology for 14 years. Nat. Med. 14:507-509.

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Nikkhah, G., Olsson, M., Eberhard, J., Bentlage, C., Cunningham, M.G., and Bjorklund, A. 1994. A microtransplantation approach for cell suspension grafting in the rat Parkinson model: A detailed account of the methodology. Neuroscience 63:57-72.

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Nikkhah, G., Rosenthal, C., Falkenstein, G., Roedter, A., Papazoglou, A., and Brandis, A. 2009. Microtransplantation of dopaminergic cell suspensions: Further characterization and optimization of grafting parameters. Cell Transplant. 18:119-133.

Roussa, E., Wiehle, M., D¨unker, N., Becker-Katins, S., Oehlke, O., and Krieglstein, K. 2006. Transforming growth factor beta is required for differentiation of mouse mesencephalic progenitors into dopaminergic neurons in vitro and in vivo: Ectopic induction in dorsal mesencephalon. Stem Cells 24:2120-2129.

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2D.5.21 Current Protocols in Stem Cell Biology

Supplement 11

Isolation of Neural Stem Cells from Neural Tissues Using the Neurosphere Technique

UNIT 2D.6

Daniela Ferrari,1 Elena Binda,1 Lidia De Filippis,1 and Angelo Luigi Vescovi1 1

Department of Biotechnology and Biosciences, University Milan-Bicocca, Milan, Italy

ABSTRACT This unit describes protocols for the derivation, characterization, and expansion of neural stem cell (NSC) lines from the adult mouse subvetricular zone (mNSCs), embryonic mouse brain and from the human fetal brain (hNSCs). NSCs can be isolated by enzymatic digestion of specific regions (NSCs niches) of the central nervous system (CNS) and grown in suspension. By using this methodology, NSCs form spherical clusters called neuropsheres, which are mechanically dissociated to a single-cell suspension and replated in the selective culture medium. Removal of growth factors and plating cells on an adherent substrate allows cells to differentiate into neurons, astrocytes, and oligodendrocytes, the main cell type of the CNS. Correct culturing of NSCs, according to this methodology, will allow cells to expand over 100 passages without alteration of cell karyotype, growth ability, and differentiation potential. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 15:2D.6.1-2D.6.18.  Keywords: neural stem cells (NSC) r subventricular zone (SVZ) r neurospheres r clonal analysis

INTRODUCTION In 1992, neural stem cells (NSCs) were identified for the first time and isolated from the subventricualr zone (SVZ) of adult mammalian brain (Reynolds and Weiss, 1992; Gritti et al., 1996; Weiss et al., 1996a,b). NSCs are multipotential precursors that grow and self-renew in culture for extensive periods of time as neurospheres, while retaining a stable capacity to generate mature, functional brain cells. NSCs represent a useful in vitro model for the study of neural developmental mechanisms as well as neuronal/glial survival and differentiation. This technique is particularly important for human studies, since NSCs from human fetuses, derived from spontaneous abortions, provide an ideal source of cells for experimental and clinical studies on cell therapy for neurodegenerative diseases. Thus far, NSC lines have been derived from the hippocampal dentate gyrus, the olfactory bulb, the SVZ surrounding the ventricles, the subcallosal zone underlying the corpus callosum, and the spinal cord of the embryonic, neonatal, and adult rodent CNS. They have been propagated in vitro using mitogens (Gritti et al., 1995, 1996, 1999, 2002; Craig et al., 1996; Reynolds and Weiss, 1996; Weiss et al., 1996a,b; Seri et al., 2006; for a review, see Bottai et al., 2003) and propagating genes (Ryder et al., 1990; Sah et al., 1997; Flax et al., 1998; Villa et al., 2000; De Filippis et al., 2007, 2008). NSC lines have also been isolated from human developing brains (Vescovi et al., 1999a,b) and adult brains (Sanai et al., 2004). A methodology that allows the isolation and expansion of NSCs using medium supplemented with growth factors (EGF and FGF2), without genetic modification, and without Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.6.1-2D.6.18 Published online November 2010 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc02d06s15 C 2010 John Wiley & Sons, Inc. Copyright 

2D.6.1 Supplement 15

tissue dissociation

+ EGF, FGF2 neurospheres (primary, secondary, tertiary, etc.)

dissociation + EGF, FGF2

no EGF, FGF2

stem cells

differentiation

Figure 2D.6.1 A schematic showing isolation of NSCs from the SVZ region of the adult mouse brain. The tissue is enzymatically digested and the single-cell suspension is exposed to EFG and FGF-2. After a period of latency ranging from 10 to 30 days in culture, free-floating clones named neurospheres can be detected and replated in the same culture conditions giving rise to secondary neurospheres. NSC can be induced to differentiate into the three CNS lineages upon growth factor removal.

serum (Fig. 2D.6.1) is described here. Methods to verify that isolated NSCs fulfill the cardinal requirements for “stemness”: proliferation ability, self-renewal capacity, functional stability, and multipotentiality (Gritti et al., 1995, 1996, 1999) are also described. This method is also known as the neurosphere assay (NSA). Since NSC lines isolated from embryonic, fetal, and adult brain display different growth requirements, and because rodent and human stem cells possess different functional characteristics, notes at each step have been included to adapt the standard protocol for the specific tissue of origin. BASIC PROTOCOL 1

ISOLATION OF NEURAL STEM CELLS FROM ADULT MOUSE SUBVENTRICULAR ZONE This protocol describes the method to isolate and expand NSCs from nervous tissue. The culture conditions are selective for the neural stem population, while most of the primary differentiated CNS cells, which are also contained in the freshly dissociated tissue, are eliminated after initial amplification steps. Four conditions should be satisfied for the NSCs to become the main cell type in these cultures: (1) low cell density (∼104 cells/cm2 ); (2) absence of serum; (3) addition of the appropriate growth factors (EGF and FGF2); and (4) absence of an adhesive substrate. This protocol focuses on isolation of adult mNSC from the SVZ of the lateral ventricles, which contains the most characterized and largest NSC niche in the adult brain, and how to establish continuous, NSC lines by growth factor stimulation.

Isolation of Neural Stem Cells from Neural Tissues

NSC can also be isolated from different CNS regions of an embryonic rodent (Reynolds and Weiss, 1992; Davis and Temple, 1994; Kilpatrick and Bartlett, 1995; Qian et al.,

2D.6.2 Supplement 15

Current Protocols in Stem Cell Biology

1997), and human fetal brain (Vescovi et al., 1999a,b) at different stages of development. At each protocol step there are specific adjustments to isolate NSC from embryonic mouse and human fetal brain.

Materials Mice (2 to 8 months old) 70% ethanol Pg solution (see recipe), ice cold Eagle’s basic salt solution (EBSS), without calcium and magnesium, ice cold Growth medium (see recipe) Papain (Worthington DBA) L-Cysteine 0.48 mM EDTA, pH 7.4 Animal anesthetic (see recipe) 10× phosphate-buffered saline (PBS), without calcium and magnesium Gentamicin Trypan blue Dissecting tools: For brain removal: large scissors, small pointed scissors, large forceps, and small spatula For SVZ dissection: scalpel, fine-curved forceps, fine-curved microscissors 15- and 50-ml polypropylene conical tubes (Falcon) Bottle top filters: low protein-binding, 0.22-μm (Millipore) Dissecting microscope Petri dishes (100-mm diameter) Rocking platform at 37◦ C Hemacytometer 6-well plates (Costar) 25-, 75-, and 162-cm2 tissue culture flasks with 0.2-μm vented filter cap (Corning) 37◦ C, 5% CO2 humidified incubator Set up experiment 1. Select the correct number of mice of the correct age. Mice should be from 2 to 8 months old. SVZ region derived from a single animal is sufficient to generate an NSC line, even though initially it is possible to pool tissue from two mice. This protocol can also be used to isolate and expand embryonic NSC from mice embryos beginning with embryonic day 10 (ED 10) or for human fetus, within weeks 8 to 12 post-conception, for routine spontaneous legal abortion. For mouse embryos, two to three animals are usually required to start a bulk culture.

2. Select dissection tools and sterilize by autoclaving 30 min at 120◦ C. During dissection procedures, immerse tools in 70% ethanol to resterilize. 3. Before dissection, make sure to have prepared and ready to use the following solutions: cold Pg solution, cold EBSS, and warm growth medium at room temperature. 4. Only for adult tissue, prepare enzymatic solution as follows for 50 ml: add 50 mg of papain to a 50-ml tube. Add 10 mg of L-cysteine and 10 mg of EDTA to a separate 50-ml tube. (Store both tubes up to 1 week at 4◦ C.) To treat the SVZ regions derived for two/three mice, 10 ml is necessary. As it might be difficult to weigh out small amounts of each reagent, it is recommended to prepare at least 50 ml of solution.

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5. Roughly 30 min before starting the enzymatic treatment, activate the papain. Add 25 ml of cold EBSS to the tube containing papain and 25 ml of cold EBSS to the cystein/EDTA tube. Vortex until the solutions are clear. Mix the two solutions together and filter (0.22-μm) to sterilize. Leave 30 min at room temperature.

Remove the brain 6. Perform animal euthanasia, removal, and dissection of brain outside the laminar flow hood. Sterilize surfaces as well as the dissection microscope with 70% ethanol. 7. Inject the mouse intraperitoneally with an overdose of animal anesthetic (2 ml/50 g mouse). a. For mouse embryos: After euthanasia, grasp the skin of the abdomen with forceps and cut through the skin and fascia to open the entire peritoneal cavity and remove the uterus. Wash uterus with sterile PBS. Using small scissors, open the uterine horns and transfer embryos to a 100-mm dish containing Pg solution. b. For human fetus: Collect human brain tissue in cold culture medium containing 5 μg/ml of gentamicin, and proceed to step 12. 8. Using large scissors, cut off the head just above the cervical spinal cord region. For mouse embryos, use the same procedure but with smaller scissors, transfer the heads to a new Petri dish containing cold Pg solution and proceed to step 12.

9. Using small pointed scissors, make a medial caudal-rostral cut and remove the skin of the head. 10. Using small scissors, make a longitudinal incision at the base of the skull and continue cutting along the sagittal suture. The cut should expose at least 2/3 of the brain to ease the following steps. While performing this step, do not damage the brain, keeping scissors as close as possible to the skull and away from brain surface. Maintaining the cerebellum in place will also allow for easier brain manipulation during SVZ dissection.

11. Using curved, pointed forceps, grasp the skull of the right hemisphere at the bottom of the incision and peel it outward to expose the brain. Repeat with the left hemisphere. Usually, the skull above the bulbs remains intact. If bulbs retrieval is necessary, remove the rostral portion of the skull, performing again a longitudinal cut and removing the skull with the forceps.

Retrieve the brain 12. To retrieve the brain, turn the head of the animal upside-down and, while cutting the optic nerves, allow the brain to slip into a 100-mm Petri dish containing ∼6 ml of cold Pg solution. Keep the Petri dish on ice while retrieving additional brains. For mouse embryos, to remove embryo brains, under the microscope, hold the head with forceps and using microscissors, perform a sagittal incision on the skull beginning at the caudal edge. The skull is very thin and brain removal is easily performed with small forceps after the sagittal cut is made. Refer to a human or rodent brain atlas for details on how to dissect the specific areas. For both human and mouse embryos, mince the tissue with microscissors and proceed to step 23.

Isolation of Neural Stem Cells from Neural Tissues

13. Wash brain with fresh Pg solution poured over brains and transfer them to a new Petri dish containing fresh Pg solution.

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Dissect the SVZ of the adult lateral ventricles 14. To dissect the forebrain SVZ region, transfer the brain to a dish without Pg solution under a dissecting microscope. Position the brain flat on its ventral surface, add 1 to 2 ml of Pg solution at the base of the brain and hold it from the caudal side using fine, curved forceps. 15. Use a scalpel to make a coronal cut 6 to 8 mm from the olfactory bulbs. At this level, the corpus callosum, the anterior commisure, and the two lateral ventricles should appear as in Figure 2D.6.2A. Repeat the cut more caudally, if necessary.

16. Make a second coronal cut to dissect an ∼2-mm thick slice embodying the lateral ventricles. 17. Position the slice with the caudal side facing the microscope. Hold the slice in position with the small forceps. Using fine, curved microscissors, cut around each ventricle to isolate them from the surrounding striatal and callosal tissue (see Fig. 2D.6.2B). 18. Place dissected SVZ into a new Petri dish containing 1 ml of cold Pg solution and mince SVZ with fine microscissors. If additional brains need to be processed, the minced tissue and solution can be transferred to a 15-ml tube using a 1000-μl pipet with the tip cut to enlarge the hole. Keep the tube on ice until enzymatic digestion.

19. Move the samples into tissue culture laminar flow hood. From this point on, use aseptic technique.

Dissociate adult brain tissue 20. Prepare one 15-ml tube with 10 ml of aerated papain solution per sample. This volume of papain solution and volumes indicated below are referred to SVZ pieces derived from 1 or 2 brains. Do not pool tissue derived from more than 2 mice.

21. With a 1000-μl pipet and 1000-μl cut tip, transfer the minced tissue pieces in Pg solution into the 15-ml tubes containing the papain solution. 22. Transfer the tubes to a rocking platform. Incubate 45 min at 37◦ C.

A

B Cc Cc Lv

Lv ac

ac

rostral side

caudal side

Figure 2D.6.2 The SVZ region to be dissected is contained in a brain slide of 2 to 3 mm, located between Bregma 1.32 (as schematically drawn in A) and Bregma 0 (as schematically drawn in B). Upon the initial cut, performed at 6 to 8 mm from the point of olfactory bulb emergence, the Cc, ac, and Lv should appear as in A. Upon the second cut, performed 2 to 3 mm rostrally, the caudal side of the brain slice should appear as in B. Use the Cc, ac, and Lv as reference brain structures. The boxed-in areas of B show the region containing the Lv area to dissect. Abbreviations: Cc, corpus callosum; ac, anterior commisure; Lv, lateral ventricle. Somatic Stem Cells

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Collect cells 23. At the end of the enzymatic incubation transfer to 15-ml tubes, and pellet tissues by centrifuging 10 min at 192 × g, room temperature. 24. Remove almost all of the supernatant overlaying the pellet. Do not use the vacuum in this step as pellet could be easily removed.

25. Add EBSS to a final 900-μl volume and dissociate the tissue with a 1000-μl pipet by forcing it through a 1-ml tip several times. Triturate the tissue until no undissociated pieces are left of the pellet. About 30 to 40 triturations should be enough to correctly dissociate a well-digested tissue. Rinse the pipet several times with medium before every dissociation step, to prevent tissue and cells from sticking to the tip walls. Avoid bubble formation. For human tissue, perform first dissociation using a glass Pasteur pipet and repeat steps 23 to 25 a second time using a 1000-μl pipet.

26. Add 5 ml of EBSS and pellet cells by centrifuging 10 min at 192 × g, room temperature. 27. Remove the supernatant leaving behind ∼200 μl. Using a 200-μl pipet with the volume set at 190 μl, gently dissociate the pellet by trituration 20 to 25 times. 28. Add 5 ml of EBSS and pellet cells by centrifuging 15 min at 192 × g, room temperature.

Count and plate cells 29. Discard supernatant and resuspend cells in 0.5 ml growth medium. 30. Dilute a 10-μl aliquot from each sample in 10 μl of 0.5% trypan blue and count viable cells using a hemacytometer (UNIT 1C.3). Adjust cell density until cells are countable on a hemacytometer. Initially try a 1:2 cells/dye dilution.

31. Seed cells at a density of 104 viable cells/cm2. in growth medium, in untreated 6-well tissue culture plates (2-ml volume) or 25-cm2 tissue culture flasks (6-ml volume). Counting cells is sometimes difficult due to the presence of debris and to the small number of cells that can be isolated. As debris is usually eliminated after a couple of passages, it is also possible to avoid cell counting at this step and culture cell suspension derived from two mice in four wells of a 6-well tissue culture plate. This yields an approximate final cell density of ∼104 cells/cm2 . For mouse embryos, the approximate number of cells yielded by this protocol using each single E14 mouse brain is: striatum, 5 × 105 cells; cortex, 2.5 × 106 cells; thalamus, 106 cells; mesencephalon, 105 cells; and spinal cord, 3 × 105 cells.

32. Incubate in a 37◦ C, 5% CO2 humidified incubator. Cells should proliferate to form spherical clusters, which eventually lift off as they grow larger. These primary spheres should be ready for sub-culturing 4 to 10 days after initial plating.

Isolation of Neural Stem Cells from Neural Tissues

Each neurosphere is derived from proliferation of NSCs and early neural progenitors and contains stem cells, differentiating progenitors, and even terminally differentiated neurons and glia. At this step of the culture, other differentiating/differentiated cells are present in the flask; these rapidly die through subculturing, while NSCs continue to proliferate, giving rise to secondary spheres that can then be further subcultured (see Basic Protocol 2). Subculturing according to the protocol described here results in a progressive enrichment of NSC in the culture.

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NEUROSPHERE CULTURE PROPAGATION: SUBCULTURING This is a protocol for successfully subculturing primary neurospheres obtained from disaggregation and culturing of NSC-containing regions of adult and embryonic mouse brain and fetal human brain. At each subculturing step, neurospheres are mechanically dissociated to a single-cell suspension and replated in the specific culture medium to generate secondary and tertiary neurospheres. If NSCs have been correctly isolated and subcultured in medium containing FGF-2 and EGF, cell number increases from two to seven fold every 4 to 10 days, depending on region, species, and developmental stage of the tissue of origin.

BASIC PROTOCOL 2

Materials Neurosphere cultures (see Basic Protocol 1) Growth medium (see recipe) 15-ml polypropylene conical tubes (Falcon) 10-ml sterile plastic pipets 25-, 75-, and 162-cm2 flasks with 0.2-μm vented filter cap (Corning) Additional reagents and equipment for trypan blue exclusion (UNIT 1C.3) 1. Subculture the neurospheres when they reach roughly 150 to 200 mm in diameter (Fig. 2D.6.3B). This will require ∼3 to 5 days for adult murine NSC lines. This step will require 3 days for embryonic mouse and 7 to 10 days for human cells. Do not dissociate neurospheres when they are too small (the yield will be low; Fig. 2D.6.3A) or allow the neurospheres to overgrow (dark region in the core of the sphere in Fig. 2D.6.3C). In this case, the number of dead cells inside the spheres will be high, trituration will be difficult, and viability of the culture will be very low.

2. Tap sides of flasks to be passaged to dislodge spheres and transfer content of the flask to 15-ml sterile plastic conical tubes using a 10-ml sterile plastic pipet. Use 5-ml fresh growth medium to rinse flask and add rinse to the tube. 3. Pellet cell suspension by centrifuging 10 min at 192 × g, room temperature. 4. Remove the supernatant leaving behind ∼250 μl.

A

B

C

Figure 2D.6.3 Neurospheres derived from NSCs cultured as described in Basic Protocols 1 and 2. (A) shows a field containing neurospheres that are too small (arrows) and ready-to-bedissociated spheres (arrowhead). (B) Neurospheres that have reached the proper size and should be dissociated for subculturing, differentiation, or collected for cryopreservation. (C) Overgrown neurospheres; note the dark region at the core of the spheres. Spheres can be dissociated but the dissociation will be more difficult to perform and at the following steps debris will be present in the culture. Scale bar = 50 μm.

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5. Use a 200-μl pipet set at a volume of 190 μl. Rinse tip with medium to avoid cell sticking inside the tip. Slightly press tip against the bottom of the tube and gently triturate pellet. Rinse down sides of tube periodically to dislodge undissociated spheres. Optimal dissociation should be obtained with 40 to 60 passages through the tip for adult rodent NSC cultures. Neurosphere dissociation is a critical step in culturing NSCs; harsh dissociation might harm stem cells and reduce stem cell number in the culture as well as leaving undissociated neurospheres. The number of times cells pass through the tip and the pressure of the tip against the bottom of the tube are two critical factors that need to be adjusted to maintain an NSC line. Neurospheres derived from mouse embryos and human fetus might require different numbers of passages through the pipet tip: 25 to 30 times for embryonic rodent cells and up to 120 times for human cells.

6. If debris or dead cells are present in excess in the culture (especially for initial passages), add 5 ml of growth medium to the dissociated pellet and centrifuge one additional time for 15 min at 192 × g, room temperature. 7. Remove supernatant, gently dissociate 10 to 20 times to desegregate pellet. 8. Count viable cells by trypan blue exclusion (UNIT 1C.3) and seed cells at 104 cells/cm2 in growth medium in untreated tissue flasks. The total cell number should increase by a factor of 2 to 5 at each passage for adult murine cells, depending on both the culture conditions and the region of origin of the cells. Viability after dissociation should never fall below 50% to 60% of the total cells. Embryonic murine cultures should increase two to ten fold and human cells two fold per passage.

9. Feed cells with fresh growth medium, i.e., replace ∼25% to 50% of the total volume in the flask, every 3 to 5 days. BASIC PROTOCOL 3

TESTING FOR MULTIPOTENCY: DEFAULT DIFFERENTIATION OF NEURAL STEM CELLS The capacity to generate all the cell lineages that belong to the tissue of origin is a fundamental property of stem cells. This protocol is a valid method to assess the multipotency of the NSC culture derived from SVZ tissue, as well as from embryonic and fetal CNS tissue. Providing the NSC with an adhesive substrate and removing growth factors according to the following methodology are sufficient to promote spontaneous differentiation of NSCs into neurons, astrocytes, and oligodendrocytes (the three major CNS cells types). These cell types can be detected by immunostaining 1 week after plating. Specific adjustments for embryonic and human NSCs are provided at each step. Multipotency should be maintained at each subculturing step if NSCs are correctly cultured and differentiated.

Materials

Isolation of Neural Stem Cells from Neural Tissues

70% ethanol Cultrex, growth factors–reduced (Trevigen) Laminin (1 mg/2 ml; Roche) DMEM, high-glucose with L-glutamine, without sodium bicarbonate or sodium pyruvate

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Neurosphere cultures (see Basic Protocol 1) NeuroCult NSC basal medium (StemCell Technologies) Complete NeuroCult NSC differentiation medium (see recipe) Glass coverslips (12- and 10-mm diameter) Glass Petri dish (11 cm2 ) 250◦ C oven Pasteur pipets Vacuum 24- and 48-well multi-well plates 37◦ C humidified incubator 15-ml polypropylene conical tubes (Falcon) Additional reagents and equipment for trypan blue exclusion (UNIT 1C.3) Set up for differentiation experiments 1. Wash glass coverslips in 70% ethanol. Dry with adsorbent paper, place in a glass Petri dish, and sterilize in a preheated oven 2 hr at 250◦ C. 2. Using a sterile Pasteur pipet connected to a vacuum, add one coverslip to each well of a 48- or 24-well plate. 3. Prepare adhesive substrate solutions as follows: a. Cultrex: 1/50 of the stock solution in cold 1× DMEM b. Laminin: 2 ml into 50 ml of 1× DMEM 4. Add 125 μl (for a 48-well plate) or 250 μl (for a 24-well plate) of the chosen adhesive substrate to the wells of the multi-well plate. 5. Incubate plates for at least 2 hr at 37◦ C in a humidified incubator to equilibrate.

Plate cells for differentiation 6. Collect neurospheres in a 15-ml tube and centrifuge cells 10 min at 192 × g, room temperature. 7. To wash cells free of growth factors, remove supernatant and resuspend cells in 10 ml of NeuroCult NSC basal medium. 8. Centrifuge 10 min at 192 × g, room temperature. 9. Remove supernatant and triturate neurospheres to a single-cell solution as described in Basic Protocol 2, steps 4 to 6. 10. Resuspend pellet in 0.5 ml of complete Neurocult differentiation medium. Count viable cells by trypan blue exclusion (UNIT 1C.3). 11. Resuspend cells in the appropriate volume of complete Neurocult differentiation medium so that the final number of cells to be plated is contained in 0.25 ml or 0.5 ml for 48- and 24-well plates, respectively. For rodent embryonic or adult cells, seed 50,000 to 70,000 cells/cm2 . For human cells, seed at 100,000 cells/cm2 . Depending on the number of cells plated, the medium may or may not have to be changed during the differentiation procedure. Plates should be checked daily. If the medium becomes acidic, it should be changed by removing ∼50% of the medium and replacing it with fresh complete Neurocult differentiation medium. Somatic Stem Cells

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12. Incubate in a 37◦ C humidified incubator. Neuronal cells can be detected as early as 2 to 4 days after plating. Under these conditions, simultaneous detection of the three cell phenotypes is usually successful at 7 days after plating. By this time point, SVZ-derived NSCs generate 5% to 8% neurons, 10% to 15% oligodendrocytes, and 70% to 80% astrocytes. Cultures derived from different regions (olfactory bulbs, rostral extension of the rostral migratory stream, dentate gyrus) or from different species generate different proportions of each cell type. Quantification of cell types in the culture can be assessed with immunocytochemical techniques using antibodies directed against β-tubulin (for neurons; Covance), glial fibrillary acidic protein (for astrocytes; Dako), and galactocerebroside-C (for oligodendrocytes;Chemicon). Human cells have a prolonged differentiation timing and to detect all cell types, the cells should be maintained in differentiation conditions for at least 15 days. BASIC PROTOCOL 4

CLONAL ANALYSIS OF NEURAL STEM CELL LINES: LIMITING DILUTION Clonal analysis and serial subcloning are likely the most difficult and time-consuming task in neural stem cell culturing. Yet, these analyses are essential to demonstrate the self-renewal capacity and the multipotency (i.e., the ability to generate neurons and both glial cell types) of the candidate cell line and to prove its “stem cell” nature. In practical terms, if the progeny of an individual clone founder cell contains cells that give rise to neurons and glia and, more importantly, contains one or more cells identical to itself (i.e., able to proliferate and produce multipotent progeny), it can be concluded that the founder cell displays stem cell features. Alternative techniques are available to perform clonal analysis (see Alternate Protocols 1 and 2). Three of these, all of which are based on the direct observation of the clone formation starting from an individual cell, are described here.

Materials Growth medium (see recipe) Neurosphere cultures (see Basic Protocol 1) Humidified chamber: glass or plastic chamber with wet gauze 96-well plates (Costar) 37◦ C humidified incubator Inverted microscope with photographic capabilities Laminin- or cultrex-coated coverslips Additional reagents and equipment for Trypan blue viable cell counting (UNIT 1C.3) 1. Warm growth medium to room temperature. 2. Prepare a humidified chamber to hold 96-well plates. 3. Collect and mechanically dissociate neurospheres to a single-cell solution (see Basic Protocol 2, steps 1 through 6). 4. Remove supernatant and resuspend pellet in 0.5 ml of growth medium. Dilute a 10-μl aliquot in 10 μl of 0.5% trypan blue and count viable cells using a hemacytometer (UNIT 1C.3).

Isolation of Neural Stem Cells from Neural Tissues

5. Resuspend cells in growth medium at a cell density of 5 to 10 cells/ml. Use a dispenser to add 100 μl of this cell suspension to each well of a 96-well plate, frequently resuspending the starting cell solution. To prepare three 96 well-plates, 300 cells/30 ml of culture medium will be needed.

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B

A

B

D

C

C

E

F

Figure 2D.6.4 Cloning of multipotent NSCs that generate neurons, astrocytes, and oligodendrocytes. A single NSC from passage 15 is shown 1 day after plating in growth medium (A) using the limiting dilution technique. This cell proliferates and by 20 divisions gives rise to a spherical clone (B-F). Single clonal spheres can be subcloned to generate secondary and tertiary spheres.

6. Incubate cells in a 37◦ C humidified incubator. 7. Carefully score plates under an inverted microscope to unequivocally identify and mark wells containing single cells. Photograph each single cell if necessary (Fig. 2D.6.4A). Make sure to use high magnification to assess that a cell is positively a single cell. Wells containing two cells or more should not be further considered for the clonal analysis.

8. Score the plate once a week; make sure the pH of the medium does not change excessively. Photograph as needed to follow the fates of single cells over time. Many of the cells will die and some will differentiate. Only a small percentage will proliferate to form a clonal sphere, which could undergo further subcloning (Fig. 2D.6.4BF). This will require 10 to 30 days, depending both on the type of cells and culture conditions.

9. Differentiate cells/neurospheres by transferring the sphere onto a laminin- or cultrexcoated coverslip and proceed as described in Basic Protocol 3. Alternatively, perform serial subcloning as described in the methylcellulose assay (see Alternate Protocol 1) or subcloning procedure (see Alternate Protocol 2).

CLONAL ANALYSIS OF NEURAL STEM CELLS: METHYLCELLULOSE ASSAY

ALTERNATE PROTOCOL 1

In this assay, neurospheres are dissociated and diluted into methylcellulose and plated. The methylcellulose prevents the movement of cells so clonal growth can be detected.

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Additional Materials (also see Basic Protocol 4) Methylcellulose powder, premium grade NeuroCult NSC basal medium (StemCell Technologies) 5-ml syringe 60-mm Petri dishes Prepare reagents 1. Warm growth medium to room temperature. 2. Prepare a humidified chamber (a glass-or plastic chamber with wet gauze on bottom) to hold 60-mm dishes. 3. Prepare a 4% (w/v) methylcellulose gel matrix in NeuroCult NSC basal medium.

Collect cells 4. Collect and dissociate neurospheres to a single-cell solution (see Basic Protocol 2, steps 1 to 6). 5. Remove supernatant and resuspend pellet in 0.5 ml of growth medium. Dilute a 10-μl aliquot in 10 μl of 0.5% Trypan blue and count viable cells using a hemacytometer (UNIT 1C.3). Make sure that the vast majority of the cells are single cells by withdrawing an aliquot and checking it under the microscope. Repeat dissociation if necessary, until only single cells are present in the suspension.

Plate cells 6. Resuspend cells in growth medium to a final cell concentration of <200 cells/ml. 7. Aspirate 2.5 ml of the cell suspension into a 5-ml syringe. 8. Aspirate 2.5 ml of the methylcellulose gel matrix into the same syringe. 9. Gently inject the mixture of cells and methylcellulose gel matrix into a 60-mm Petri dish, avoiding bubbling and foaming. 10. Using the same syringe, resuspend the mixture multiple times until a semi-solid homogeneous gel has formed and the single cells are thoroughly dispersed.

Evaluate the culture 11. The day after plating, score the plate to identify single hypertrophic (bright) cells. 12. Mark the position of these cells on the plate with a fine marker and take microphotographs over time. If the appropriate set up is available, use tissue culture flasks rather than Petri dishes, seal tightly and use time-lapse cinematography to monitor clone formation continuously.

13. When clonal spheres have been generated, perform subcloning of individual spheres by repeating the steps in this protocol or performing Alternate Protocol 2; or initiate differentiation as described in Basic Protocol 3. ALTERNATE PROTOCOL 2

CLONAL ANALYSIS OF NEURAL STEM CELLS: SUBCLONING PROCEDURE Subcloning of individual neurospheres allows for isolation of NSC from the neurosphere.

Additional Materials (also see Basic Protocol 4) Isolation of Neural Stem Cells from Neural Tissues

Neurospheres (see Basic Protocol 3 or Alternate Protocol 1) 5-ml microcentrifuge tubes 12-, 24-, or 48-well plates

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1. Transfer individual clonal spheres to 5-ml microcentrifuge tubes containing 1 ml of growth medium (1 sphere/tube) using a sterilized 200-μl pipettor set at 180 μl. Rinse the tip with medium first to avoid cells sticking to the tip wall. If you want to generate a clonal cell line, use the limiting dilution protocol and then dissociate single clonal spheres inside their own dish (96-well dish) without transferring them.

2. Centrifuge 10 min at 192 × g, room temperature. Remove supernatant leaving behind ∼200 μl of medium. 3. Using a 200-μl pipettor set at 180 μl, mechanically dissociate spheres to a single-cell suspension. 4. Plate all the cell suspension in a 48-, 24-, or 12-well plate (depending on the number of viable cells) and incubate cells in a 37◦ C humidified chamber. Embedding cells into methylcellulose prior to plating (see Alternate Protocol 1) is strongly recommended to avoid cell aggregation.

5. Within 1 hr of plating, count the number of cells obtained by dissociation of each clone under the microscope. A subset of these cells will proliferate giving rise to secondary clones.

6. Calculate the cloning efficiency, normalizing the number of secondary clones by the total number of cells in the same well, as assessed by direct observation 1 hr after dissociation. 7. Differentiate individual secondary clones to assess their multipotentiality (see Basic Protocol 3) or transfer the neurospheres to a 5-ml microcentrifuge tubes (1 sphere/tube) to undergo further subcloning. 8. If a clonal cell line has to be generated, pool secondary spheres derived from a single primary sphere, mechanically dissociate to a single-cell suspension and plate at a cell density of 104 cells/cm2 in the appropriate medium. Subculture until a bulk culture is established.

CRYOPRESERVATION OF NEUROSPHERES Cryopreservation of NSC lines in liquid nitrogen, as described in this section, does not affect NSC properties even after repeated cycles of freezing and thawing. Therefore, this protocol allows the establishment of an NSC cell bank for research or clinical purposes.

BASIC PROTOCOL 5

Materials 100% isopropanol Dimethylsulfoxide (DMSO) Growth medium (see recipe) Neurospheres Liquid nitrogen tank 70% ethanol Cryo 1◦ C freezing container (Nalgene) 15-ml tubes 2-ml cryovials 37◦ C water bath 15-ml plastic tube Flasks Somatic Stem Cells

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Freeze neurospheres 1. Allow the freezing container to come to room temperature. 2. Fill the container with isopropanol. 3. Prepare the freezing medium: 10% DMSO in growth medium (store not more than 1 week at 4◦ C). 4. Collect spheres in a 15-ml tube by gentle pipetting. Centrifuge 10 min at 192 × g, room temperature. Do not freeze neurospheres that are overgrown or too small, this will result in poor survival after thawing. Optimal neurosphere size is the same as for subculturing.

5. Discard all the supernatant and resuspend the pellet in freezing medium using a 2-ml pipet to gently disaggregate the pellet. In 2 ml of freezing medium, collect and freeze in one 2-ml cryovial neurospheres derived from a 75-cm2 flask. For neurospheres from a 175-cm2 flask, resuspend in 4 ml freezing medium and distribute into two cryovials. 6. Transfer vials into the freezing container and then into a –80◦ C freezer and allow vials to reach –80◦ C (check cooling rate from manufacturer’s instructions). 7. Transfer vials into a liquid nitrogen tank for long-term storage.

Thaw cryopreserved neurospheres 8. Warm growth medium to room temperature. 9. Transfer cryovial(s) from liquid nitrogen to a 37◦ C water bath and leave until thawed (2 to 5 min). 10. Wipe entire cryovial with 70% ethanol. 11. Slowly transfer cell suspension from cryovial to a 15-ml plastic tube containing 5 ml of warm growth medium. 12. Centrifuge cell suspension for 10 min at 192 × g, room temperature, and remove most of the supernatant. 13. Gently resuspend pellet in fresh growth medium and plate in flask(s) of appropriate size.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Animal anesthetic To prepare 200 ml of avertin solution, mix 2.5 g of 2,2,2-tribromoethanol (97% tribromoethyl alcohol; Sigma-Aldrich) and 5 ml of 2-methyl-2-butanol (tert-amyl alcohol, ≥99%, Reagent Plus; Sigma-Aldrich). Warm the solution in a 37◦ C bath and mix until the solution is clear (5 to 15 min). Add distilled water to a final volume of 200 ml, 12.5 mg/ml final concentration. Store up to 1 week at 4◦ C protected from light.

Complete differentiation medium Isolation of Neural Stem Cells from Neural Tissues

Thaw one bottle containing 50 ml of NeuroCult NSC differentiation supplements (StemCell Technologies) at room temperature or overnight at 4◦ C. Dispense supplements into 10-ml aliquots and store up to 1 month at −20◦ C.

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A white precipitate may form in the supplements during storage at −20◦ C; it will disappear when supplement is completely thawed and mixed well. However, repeated freezing and thawing of aliquots is not recommended.

Add the entire 50-ml volume of NeuroCult NSC differentiation supplements to a bottle containing 450 ml of NeuroCult NSC basal medium (StemCell Technologies; or 1 ml of differentiation supplements to every 9 ml basal medium). Prepare fresh.

Glucose, 30% (w/v) Mix 30 g glucose in 100 ml water. Filter sterilize using a 0.22-μm filter. Store up to 1 month at 4◦ C.

Growth factor stocks Reconstitute EGF (Peprotech) to 500 μg/ml and FGF2 (bFGF; Peprotech) to 100 μg/ml. Dispense into sterile tubes (40 μl for EGF and 100 μl for FGF2) and store at –20◦ C.

Growth medium Thaw one bottle containing 50 ml of NeuroCult NSC proliferation supplements (StemCell Technologies). Add the entire volume of proliferation supplements to a bottle containing 450 ml of NeuroCult NSC basal medium (StemCell Technologies). Add 20 μl EGF stock (20 ng/ml final concentration; see recipe for growth factor stocks) or/and 1000 μl FGF-2 stock (10 ng/ml final concentration; see recipe for growth factor stocks). Store up to 2 weeks at 4◦ C.

Pg solution To prepare 500 ml of Pg solution, add: 50 ml of sterile 10× phosphate-buffered saline (PBS) 5 ml of 100× penicillin/streptomycin 10 ml 30% (w/v) glucose (final concentration 0.6%; see recipe) 435 ml water Filter with a 0.2-μm filter and protect from light to avoid degradation of penicillin/streptomycin Prepare fresh COMMENTARY Background Information Since their identification and upon the establishment of NSC lines from different regions of embryonic, fetal, and adult brain (see Bottai et al., 2003, for review), NSCs have been revealed to be a useful tool for a broad spectrum of applications, both in vitro as a model of neural development and neuronal/glial survival and differentiation (Bjornson et al., 1999; Galli et al., 2000, 2002; Cedrola et al., 2003) and in vivo as a source of cells for experimental and, recently, clinical studies for cell therapy in many neurodegenerative diseases. NSCs have been maintained in vitro using mitogens (Gritti et al., 1995, 1996, 1999, 2002; Craig et al., 1996; Reynolds and Weiss, 1996; Weiss et al., 1996a,b; Seri et al., 2006) and with propa-

gating genes (Ryder et al., 1990; Sah et al., 1997; Flax et al., 1998; Villa et al., 2000; De Filippis et al., 2007, 2008). Whereas the immortalization of NSCs enhances the rate of proliferation, survival ability, and multipotency (De Filippis et al., 2007, 2008), described here is a methodology for the isolation and expansion of NSCs using growth factors (EGF and FGF2)–containing medium without genetic modification and without serum. The method described in this unit represents a safe NSC culture method for cell therapy development. When establishing an NSC line, it is important to verify the basal stem cell properties of the isolated population. Formation of primary neurospheres is not sufficient to establish the presence of NSC into the culture. Dissociated neural tissue, especially that derived from

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embryonic and fetal brains, contains lineagerestricted progenitors, which can behave as stem cells for a limited number of passages; indeed such progenitors survive and give rise to secondary and tertiary neurospheres, leading to misinterpretation of results. A further limiting factor for NSC line isolation is cell density: if the cell density exceeds an optimal threshold (see Basic Protocols 1 and 2), aggregating clusters of cells can be erroneously considered neurospheres. Hence, serial subculturing over six to seven passages, simultaneously with retention of multipotency and self-renewal abilities, is required to establish a bona fide NSC cell line. The methodology for the isolation and expansion of NSC lines from different neural tissues (see Basic Protocols 1 and 2), as well as for the evaluation of multipotency (see Basic Protocol 3) and self-renewal (see Basic Protocol 4 and Alternate Protocols 1 and 2) is described in this unit. The term “neural stem cells” is applied to precursors that, most likely, occupy quite distinct hierarchical positions within the normal neurogenic lineage. The risk of false-positive identification of neural stem cells in a culture is very high when the neurosphere assay is applied loosely on a too short-term basis, and the formation of neurospheres is construed as indicating the presence of stem cells without the required long-term population and growth kinetic analysis. Recently, a central tenet of the neurosphere assay, that all neurospheres are derived from a stem cell and that the NSA can be used to estimate NSC frequency, has been challenged. It has been demonstrated by modeling and experimental evidence that this premise is false. In response, two new assays have been developed: (1) neural colony– forming cell assay (N-CFCA; Louis et al., 2008) and (2) a mathematical model that now provides us with a manner to meaningfully estimate NSC frequency and measure symmetric stem cell divisions, respectively (Reynolds and Rietze, 2005).

Critical Parameters and Troubleshooting

Isolation of Neural Stem Cells from Neural Tissues

Basic Protocol 1 Because different growth rates have been shown for NSC lines deriving from murine or human brain and for embryonic versus adult brain (Bottai et al., 2003), the source of neural tissue should be carefully evaluated and considered when evaluating growth rate.

Because the stem cell fraction in tissue samples is relatively low, especially for adult tissue, dissection should be carried out as quickly as possible (within 2 hr) to avoid tissue degradation processes. Over time, tissue becomes soft and sticky and may be difficult to dissect. Basic Protocol 2 As already mentioned for Basic Protocol 1, the source of tissue for NSC line establishment is a determining factor for the expansion of neurospheres, and several cautions have to be adopted for the correct neurosphere assay (NSA). The abilities of the operator when subculturing are critical for the success of NSC isolation and amplification: harvesting failure of the cells, inefficient or too harsh mechanical dissociation, and pH unbalance of the culture medium, play prominent roles in the subculturing efficiency. To avoid pitfalls in the NSA procedure, the following are some guidelines: (1) When harvesting neurospheres, tap the flask before collecting neurospheres, wash flask with fresh growth medium to be sure all neurospheres are collected, and use polypropylene tubes for centrifugations. (2) For mechanical dissociation, be sure to use the correct pipet tips, avoid foaming by setting the pipettor volume to a value of ∼20 μl lower than the actual capacity of the tip, achieve dissociation to single cells with the number of triturations indicated in the protocol and, if necessary, modify the pressure of the tip against the tube bottom. Do not leave neurospheres in pellets >1 hr before processing. Choose the correct stage of neurosphere development (see Fig. 2D.6.3) for neurosphere dissociation. (3) Always keep the bottle of culture medium well capped and limit exposure to air CO2 when working. Do not use medium >1 month after preparation.

Anticipated Results This protocol is used to establish karyotypically normal NSC lines that are stable regarding their growth kinetics, self renewal, and differentiation potential. For adult mouse SVZ NSCs, each subculturing step results in a twoto five-fold increase in cell number (up to tenfold for mouse embryonic NSC and two-fold for human fetal NSCs). As the initial number of NSCs in the dissociated tissue may depend on many variables as described above, a minimum of 106 cells (for mouse tissue) and 5 × 105 cells (for human tissue) is expected 2 weeks after the initial tissue dissociation.

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Time Considerations Basic Protocol 1 It takes 40 min to isolate SVZ regions from three animals or one region from embryos deriving from each pregnant mouse. Enzymatic digestion of tissue takes 30 to 40 min. Its requires 30 min to set up an NSC primary culture. Basic Protocol 2 It takes 15 min to subculture each flask. To establish a stable NSC line from adult mouse brain, 1 month of passaging is required, 20 days from embryonic mouse brain, 2 months from human fetal brain. Basic Protocol 3 For adhesive substrate preparation, 2 hr are required and 30 min are required to prepare dissociated neurospheres and plate a complete 24/48-well plate. Basic Protocol 4 To prepare a complete 96-well plate, 30 min are required. Alternate Protocol 1 It takes 20 min to prepare methyl-cellulose gel matrix, 30 min for cell harvesting and counting, and 10 min for seeding a petri dish. Alternate Protocol 2 Thirty minutes are required for subcloning at each passage. Basic Protocol 5 It takes 15 min to collect neurospheres into the cryovial. It takes an additional ∼6 hr to move the cryovial to liquid nitrogen.

Literature Cited Bjornson, C.R., Rietze, R.L., Reynolds, B.A., Magli, M.C., and Vescovi, A.L. 1999. Turning brain into blood: A hematopoietic fate adopted by adult neural stem cells in vivo. Science 283:534-537. Bottai, D., Fiocco, R., Gelain, F., Defilippis, L., Galli, R., Gritti, A., and Vescovi, L.A. 2003. Neural stem cells in the adult nervous system. J. Hematother. Stem Cell Res. 12:655-670. Cedrola, S., Guzzi, G., Ferrari, D., Gritti, A., Vescovi, A.L., Pendergrass, J.C., and La Porta, C.A. 2003. Inorganic mercury changes the fate of murine CNS stem cells. FASEB J. 17:869871. Craig, C.G., Tropepe, V., Morshead, C.M., Reynolds, B.A., Weiss, S., and van der Kooy, D. 1996. In vivo growth factor expansion of endogenous subependymal neural precursor cell

populations in the adult mouse brain. J. Neurosci. 16:2649-2658. Davis, A.A. and Temple, S. 1994. A self-renewing multipotential stem cell in embryonic rat cerebral cortex. Nature 372:263-266. De Filippis, L., Lamorte, G., Snyder, E.Y., Malgaroli, A., and Vescovi, A.L. 2007. A novel, immortal, and multipotent human neural stem cell line generating functional neurons and oligodendrocytes. Stem Cells 25:2312-2321. De Filippis, L., Ferrari, D., Rota Nodari, L., Amati, B., Snyder, E., and Vescovi, A.L. 2008. Immortalization of human neural stem cells with the c-myc mutant T58A. PLoS ONE 3:e3310. Flax, J.D., Aurora, S., Yang, C., Simonin, C., Wills, A.M., Billinghurst, L.L., Jendoubi, M., Sidman, R.L., Wolfe, J.H., Kim, S.U., and Snyder, E.Y. 1998. Engraftable human neural stem cells respond to developmental cues, replace neurons, and express foreign genes. Nat. Biotechnol. 16:1033-1039. Galli, R., Borello, U., Gritti, A., Minasi, M.G., Bjornson, C., Coletta, M., Mora, M., De Angelis, M.G., Fiocco, R., Cossu, G., and Vescovi, A.L. 2000. Skeletal myogenic potential of human and mouse neural stem cells. Nat. Neurosci. 3:986991. Galli, R., Fiocco, R., De Filippis, L., Muzio, L., Gritti, A., Mercurio, S., Broccoli, V., Pellegrini, M., Mallamaci, A., and Vescovi, A.L. 2002. Emx2 regulates the proliferation of stem cells of the adult mammalian central nervous system. Development 129:1633-1644. Gritti, A., Cova, L., Parati, E.A., Galli, R., and Vescovi, A.L. 1995. Basic fibroblast growth factor supports the proliferation of epidermal growth factor-generated neuronal precursor cells of the adult mouse CNS. Neurosci. Lett. 185:151-154. Gritti, A., Parati, E.A., Cova, L., Frolichsthal, P., Galli, R., Wanke, E., Faravelli, L., Morassutti, D.J., Roisen, F., Nickel, D.D., and Vescovi, A.L. 1996. Multipotential stem cells from the adult mouse brain proliferate and self-renew in response to basic fibroblast growth factor. J. Neurosci. 16:1091-1100. Gritti, A., Frolichsthal-Schoeller, P., Galli, R., Parati, E.A., Cova, L., Pagano, S.F., Bjornson, C.R., and Vescovi, A.L. 1999. Epidermal and fibroblast growth factors behave as mitogenic regulators for a single multipotent stem celllike population from the subventricular region of the adult mouse forebrain. J. Neurosci. 19:32873297. Gritti, A., Bonfanti, L., Doetsch, F., Caille, I., Alvarez-Buylla, A., Lim, D.A., Galli, R., Verdugo, J.M., Herrera, D.G., and Vescovi, A.L. 2002. Multipotent neural stem cells reside into the rostral extension and olfactory bulb of adult rodents. J. Neurosci. 22:437445. Kilpatrick, T.J. and Bartlett, P.F. 1995. Cloned multipotential precursors from the mouse cerebrum require FGF-2, whereas glial restricted

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precursors are stimulated with either FGF-2 or EGF. J. Neurosci. 15:3653-3661.

adult human brain contains neural stem cells but lacks chain migration. Nature 427:740-744.

Louis, S.A., Rietze, R.L., Deleyrolle, L., Wagey, R.E., Thomas, T.E., Eaves, A.C., and Reynolds, B.A. 2008. Enumeration of neural stem and progenitor cells in the neural colony-forming cell assay. Stem Cells 26:988-996.

Seri, B., Herrera, D.G., Gritti, A., Ferron, S., Collado, L., Vescovi, A., Garcia-Verdugo, J.M., and Alvarez-Buylla, A. 2006. Composition and organization of the SCZ: A large germinal layer containing neural stem cells in the adult mammalian brain. Cereb. Cortex 16:i103-i111.

Qian, X., Davis, A.A., Goderie, S.K., and Temple, S. 1997. FGF2 concentration regulates the generation of neurons and glia from multipotent cortical stem cells. Neuron 18:81-93. Reynolds, B.A. and Weiss, S. 1992. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707-1710. Reynolds, B.A. and Weiss, S. 1996. Clonal and population analyses demonstrate that an EGFresponsive mammalian embryonic CNS precursor is a stem cell. Dev. Biol. 175:1-13. Reynolds, B.A. and Rietze, R.L. 2005. Neural stem cells and neurospheres—Re-evaluating the relationship. Nat. Methods 2:333-336. Ryder, E.F., Snyder, E.Y., and Cepko, C.L. 1990. Establishment and characterization of multipotent neural cell lines using retrovirus vector-mediated oncogene transfer. J. Neurobiol. 21:356-375. Sah, D.W., Ray, J., and Gage, F.H. 1997. Bipotent progenitor cell lines from the human CNS. Nat. Biotechnol. 15:574-580. Sanai, N., Tramontin, A.D., Quinones-Hinojosa, A., Barbaro, N.M., Gupta, N., Kunwar, S., Lawton, M.T., McDermott, M.W., Parsa, A.T., ManuelGarcia Verdugo, J., Berger, M.S., and AlvarezBuylla, A. 2004. Unique astrocyte ribbon in

Vescovi, A.L., Gritti, A., Galli, R., and Parati, E.A. 1999a. Isolation and intracerebral grafting of nontransformed multipotential embryonic human CNS stem cells. J. Neurotrauma 16:689693. Vescovi, A.L., Parati, E.A., Gritti, A., Poulin, P., Ferrario, M., Wanke, E., Frolichsthal-Schoeller, P., Cova, L., Arcellana-Panlilio, M., Colombo, A., and Galli, R. 1999b. Isolation and cloning of multipotential stem cells from the embryonic human CNS and establishment of transplantable human neural stem cell lines by epigenetic stimulation. Exp. Neurol. 156:71-83. Villa, A., Snyder, E.Y., Vescovi, A., and MartinezSerrano, A. 2000. Establishment and properties of a growth factor-dependent, perpetual neural stem cell line from the human CNS. Exp. Neurol. 161:67-84. Weiss, S., Dunne, C., Hewson, J., Wohl, C., Wheatley, M., Peterson, A.C., and Reynolds, B.A. 1996a. Multipotent CNS stem cells are present in the adult mammalian spinal cord and ventricular neuroaxis. J. Neurosci. 16:75997609. Weiss, S., Reynolds, B.A., Vescovi, A.L., Morshead, C., Craig, C.G., and van der Kooy, D. 1996b. Is there a neural stem cell in the mammalian forebrain? Trends Neurosci. 19:387-393.

Isolation of Neural Stem Cells from Neural Tissues

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Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

UNIT 2E.1

Yuzo Niki1 1

Ibaraki University, Ibaraki, Japan

ABSTRACT This unit describes how to collect, culture, and establish stable cell lines of ovarian somatic and germline stem cells of Drosophila. We also describe a protocol for culturing embryonic cells that overexpress growth factors, which serve as a source for conditioned C 2009 by John Wiley & Sons, medium. Curr. Protoc. Stem Cell Biol. 10:2E.1.1-2E.1.9.  Inc. Keywords: Drosophila r somatic and germline stem cells r isolation r expansion

INTRODUCTION Germline stem cells (GSCs) and their niches have been extensively studied in vivo in Drosophila. The concept that stem cells are controlled by particular microenvironments known as niches has been widely suggested by various in vivo approaches (reviewed by Fuller and Spradling, 2007). In vitro systems, however, are powerful and indispensable for analyzing the interactions between GSCs and their niches directly and biochemically (reviewed by Niki, 2008). Thus, this unit describes how to collect, culture, and establish stable cell lines of ovarian somatic stem (OSS) cells and GSCs from tumorous mutant ovaries. NOTE: The following tissue culture procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All incubations are performed at 25◦ C and do not need any special equipment such as a CO2 incubator. NOTE: Most fly laboratories add live (dry) yeast as food to raise flies. It is hard to remove or kill yeast by treating with any kind of antibiotics and antimicrobial drugs for tissue and cell cultures. Instead of live yeast, we feed “microwaved” yeast (see recipe). NOTE: Storing many ovaries at once in a depression glass slide frequently results in contamination with microorganisms. To reduce the risk of contamination, it is recommended that several ovaries from only one or two females be collected in a depression glass slide.

ISOLATION AND CULTURE OF OVARIAN SOMATIC STEM CELLS AND GERMLINE STEM CELLS

BASIC PROTOCOL

The protocol below describes a common procedure for collecting and culturing OSS cells and GSCs. To obtain sufficient numbers of OSS cells and GSCs, we used females of either the w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ] or w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ) P[vasa-egfp] genotype. These constructs have the wild-type allele of a bam gene ligated with a heat shock promoter. Germline cells are marked with ovo-lacZ and vasa-egfp. In these bam homozygous females, OSS cells and GSCs expand as the female ages. Somatic Stem Cells Current Protocols in Stem Cell Biology 2E.1.1-2E.1.9 Published online September 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02e01s10 C 2009 John Wiley & Sons, Inc. Copyright 

2E.1.1 Supplement 10

Materials 20- to 30-day-old female flies of w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ] or w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ) P[vasa-egfp] genotype (Fig. 2E.1.1A,B) 70% (v/v) ethanol Drosophila phosphate-buffered saline (PBS; Robb, 1969), sterile Culture medium (see recipe) Penicillin/streptomycin (see recipes), optional Distilled water DMSO Liquid nitrogen 15-ml conical tubes Single concave depression glass slides Forceps Sterilized tungsten needles 96-well tissue culture plates Sealed container (e.g., Tupperware) Phase-contrast and fluorescent microscope 200-μl pipet tips Cryotubes (Nunc) −20◦ and −80◦ C freezers 1.5-ml microcentrifuge tubes Isolate female GSCs (fGSCs) from bag-of-marbles (bam) ovaries 1. Sterilize 20- to 30-day-old female flies homozygous for the bam mutation, as described above, with 5 ml 70% ethanol for 10 min in a 15-ml conical tube. 2. Immerse several adult flies in a drop of sterilized PBS on a single concave depression glass slide. 3. Using forceps, dissect the ovaries carefully so as to not injure the gut. 4. Wash the dissected ovaries three times with culture medium. If necessary, add a mixture of penicillin and streptomycin to the culture medium. 5. Dissociate ovaries into ovarioles and then fragment each ovariole into pieces with fine tungsten needles. 6. Remove any membranous debris carefully. 7. Wash the cell masses three times, each time with 100 μl culture medium.

Plate the cells 8. Dispense 100 μl of the culture medium–containing cell masses into interior wells of a 96-well tissue culture plate. 9. Add new culture medium to make a final volume of 200 μl in each well. 10. Add 200 ml of distilled water in the external marginal wells of a 96-well culture plate to reduce the evaporation of the culture medium. 11. Store the culture plates in a sealed container, such as a Tupperware container, at 25◦ C. Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

12. Check the cells with a phase-contrast microscope daily and exchange half of the culture medium every week.

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A

B

C

D

E

Figure 2E.1.1 (A) A pair of tumorous bam ovaries from a 30-day-old female Drosophila melanogaster flies. (B) A living bam ovariole showing vasa-gfp positive GSCs. (C) A stable line named the fGS/OSS line. Female GSCs are located on the somatic cells that originated from follicle stem cells. (D) Cellular clump formed after confluence. (E) Living OSS cells stained with Hoechst 33248. (F) OSS cells in the subconfluent condition. Bar = 50 μm.

13. Scrape cellular clumps with a 200-μl pipet tip and disperse them by gentle pipetting. Split the cells 1:2 or 1:4 into new wells of the same culture plate when they become confluent. For expanding cells, it is much better to transfer cells into new wells than to leave them in old wells. Once split, cells continue to divide and expand rapidly thereafter. It takes ∼20 hr for the doubling of cell number. Split cultured cells before they become confluent. Note that GSCs are sensitive to cell density and sometimes disappear rapidly if you leave the cells alone after they have become confluent (Fig. 2E.1.1D). During primary culture, the cells start to spread and continue to divide for 1 to 2 weeks after initiation of culture, but cease to divide thereafter. It is important to continue to exchange the medium once every week even if the cultured cells have ceased to grow and turned brown or black, a symptom of cell death. There will be cases in which several transparent cells appear at the periphery of the colored cellular aggregates and begin to divide again one or several months after culture. These somatic cells originate from

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daughter cells of the somatic stem cells. The cells then start expanding dramatically and cover the bottom of the culture well to form a sheet. There will also be cases in which round, large cells, characteristic of GSCs, appear in the somatic sheet. GSCs associated with somatic cells continue to divide as the somatic sheet expands (Fig. 2E.1.1C).

Establish cell lines consisting of only ovarian somatic cells 14. Seed 1 to 50 cells, from the above GSC/OSS cell culture, in each well of a 96-well plate separately, after dissociating with a 200-μl tip. 15. Select wells consisting only of somatic cells by checking the vasa-EGFP activity in the live condition with a phase-contrast and fluorescent microscope. 16. Repeat the dilution and expansion of the somatic cells several times until the somatic cells show the same phenotype in all subpopulations (Fig. 2E.1.1E,F).

Freeze and thaw cultured cells 17. Add 20 μl DMSO to the well and gently disperse the cells by pipetting. Transfer 1–10 × 104 cells into a cryotube. 18. Store the cells overnight at −20◦ C. 19. Transfer the cells to a −80◦ C deep freezer and then store them in liquid nitrogen. 20. Transfer the cryotube to a clean bench and warm the tubes with water at 25◦ C to thaw the frozen cells. 21. Wash the cells with 400 μl culture medium, transfer them into a 1.5-ml microcentrifuge tube, and remove the DMSO by centrifuging 1 min at 500 × g in a microcentrifuge at 25◦ C. 22. Seed 1–10 × 103 cells into the wells of a culture plate. 23. Add fresh culture medium to make the final volume 200 μl in each well.

Phenotype the cells 24. Discriminate GSCs and OSS cells from each other by size and morphology: GSCs are round and 10-μm in diameter and are usually located on the OSS cell sheet (Fig. 2E.1.1C), whereas OSS cells are small and flattened and are <6-μm in diameter, like epithelial cells (Fig. 2E.1.1F). GSCs are positive for any of the GSC markers— Vasa, Nanos, and dot-shaped fusome. OSS cells are positive for FasIII, a marker of follicle stem cells and their descendant cells (Niki et al., 2006). SUPPORT PROTOCOL 1

PREPARATION OF FLY EXTRACT We prepare fly extract (FE) according to the method of Currie et al. (1988).

Materials Young adult flies, 2 to 3 days after emergence 70% (v/v) ethanol Drosophila phosphate-buffered saline (PBS; Robb, 1969), sterile M3 (BF) medium (see recipe; also see Cross and Sang, 1978), 25◦ C Culture medium (see recipe)

Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

15-ml conical tubes Dounce homogenate 1.5-ml microcentrifuge tubes Centrifuge

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1. Collect young adult flies within 2 to 3 days after emergence. You need 200 flies to make a 1.5-ml aliquot of FE.

2. Sterilize the flies with 5 ml 70% ethanol for 10 min in a 15-ml conical tube. 3. Wash the sterilized flies three times with 5 ml sterilized PBS and once with 5 ml M3-BF culture medium. 4. Homogenize 200 sterilized flies with 1.2 ml of culture medium with a Dounce homogenizer. Transfer the homogenate to a 1.5-ml tube. 5. Centrifuge the homogenate 20 min at 1,500 × g, 4◦ C. 6. Heat-inactivate the supernatant at 60◦ C for 5 min. 7. Centrifuge the heat-inactivated supernatant 10 min at 6,000 × g, 4◦ C. 8. Transfer the supernatant into another 1.5-ml tube. 9. Repeat the centrifugation 20 min at 10,000 × g, 4◦ C. Transfer the supernatant to another 1.5-ml tube. 10. Centrifuge the supernatant 60 min at 15,000 × g, 4◦ C again. 11. Adjust the final volume of each aliquot to 1.5 ml by adding culture medium. 12. Store the FE up to 3 monhts at −20◦ C.

PREPARATION OF CONDITIONED MEDIUM FROM EMBRYOS OVEREXPRESSING GROWTH FACTORS

SUPPORT PROTOCOL 2

We prepare embryonic primary cultures according to the method of Ui et al. (1987). Ueda et al. (2007) reported a protocol for the mass culture of flies to obtain a large number of embryonic embryos. From our experience, however, stable cell lines of embryonic cell origin can be established from eggs collected from several hundred adult flies.

Materials actin-Gal 4 Drosophila lines (National Institute of Genetics, Mishima, Japan) UAS-dpp, wingless, or hedgehog Drosophila lines (Bloomington fly stock center) 35-mm culture dish filled with 10% agar pasted with “microwaved” yeast (see recipe) Saponated cresol solution (Japanese Pharmacopoeia) 2.5% sodium hypochlorite (NaOCl) 0.1% Triton X100/PBS (PBT) Culture medium (see recipe) 15-ml conical tube with bottom cut and stainless steel mesh attached 1.5-ml microcentrifuge tubes Microhomogenizer 100- to 150-μm nylon mesh 48- or 96-well culture plates 15-ml conical tubes 0.22-μm syringe filter Obtain embryos 1. Mate actin-Gal4 flies with UAS-growth factor flies to obtain offspring embryos that overexpress the growth factors. 2. Collect staged embryos on 35-mm agar dishes pasted with “microwaved” yeast.

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3. Transfer collected embryos into a 15-ml conical tube in which the bottom has been cut and stainless mesh has been attached. 4. Wash the embryos with streaming water to remove yeast. 5. Transfer the embryos into a 1.5-ml microcentrifuge tube and add 800 μl of a 1/10 diluted saponated cresol solution for 10 min to sterilize the embryos. 6. Dechorionate embryos with 800 μl 2.5% NaOCl for several minutes. Remove the cresol solution before adding the NaOCl.

7. Wash dechorionated embryos with 800 μl sterilized PBT (0.1% Triton X100 in PBS).

Prepare embryo homogenate 8. Suspend dechorionated embryos in 800 μl culture medium in a 15-ml microcentrifuge tube. 9. Homogenize the embryos with a microhomogenizer. 10. Filter the homogenized cells with a nylon mesh 100- to 150-μm in diameter. 11. Repeat suspensions and centrifuge 3 times for 1 min at 500 × g, 25◦ C. 12. Resuspend the cell pellets in 400 ml fresh culture medium.

Plate the cells 13. Adjust the cell density to 0.5–1.0 × 106 cells/ml. 14. Seed 1–10 × 104 cells into wells of 48- or 96-well culture plates. 15. Continue to culture these primary embryonic cells by exchanging half the volume with fresh culture medium once every 5 to 7 days.

Prepare conditioned medium 16. Collect the medium from the primary cultures in a 15-ml conical tube and centrifuge 1 min at 500 × g, 25◦ C, to make the conditioned medium. 17. Mix the supernatant with fresh culture medium in a 1:1 ratio in a 1.5-ml conical tube. 18. Filter-sterilize the conditioned medium with a 0.22-μm syringe filter. 19. Store the conditioned medium up to 7 days at 4◦ C. SUPPORT PROTOCOL 3

FIXATION AND STAINING OF CULTURED DROSOPHILA CELLS Cultured Drosophila cells are fixed and stained for markers of OSS and GSC.

Materials

Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

Cultures of Drosophila cells Ice 4% (w/v) paraformaldehyde in PBS 0.1% Triton X100 in PBS (PBT) Primary antibodies Rabbit anti-Vasa antibody (S. Kobayashi, National Institute for Basic Biology, and A. Nakamura, Riken Center for Developmental Biology) at a 1:250 dilution Rabbit anti-Spectrin antibody (R. Dubreuil, University of Illinois-Chicago) at a 1:1000 dilution Mouse anti-Fusome antibody (F. Maruo, Tsukuba University) at a 1:3 dilution Mouse anti-GFP antibody (Molecular Probes)

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Current Protocols in Stem Cell Biology

Mouse anti-Drosophila Dpp antibody (R&D systems, Lot number FRW02) at 1:100 to 1:200 dilutions Mouse anti-Fasciclin III (FasIII) antibody at a 1:50 dilution (Developmental Studies Hybridoma Bank, Iowa University) Secondary antibodies: FITC- or TRITC-conjugated goat anti-rabbit IgG or Alexa Fluor 546 (Molecular Probes) at 1:100 or 1:200–1:500 dilution in 5% (w/v) BSA FITC- or TRITC-conjugated goat anti-mouse IgG or Alexa Fluor 488 (Molecular Probes) at 1:100 or 1:200–1:500 dilution in 5% (w/v) BSA Hoescht 33258 or 33342 50% glycerol in PBS or Aqua Poly-Mount (Polysciences) Appropriate microscopes 1. Fix cells with 200 μl 4% formaldehyde in PBS for 5 min on ice. 2. Wash the fixed cells three times, each time with 200 μl PBT for 60 min. 3. Stain the cells with primary antibody for 60 min at room temperature. To avoid mislocalization during preparation, PBS is used instead of PBT throughout preparation when anti-Dpp antibody is used.

4. Wash the cells three times, each time with 200 μl PBT for 60 min. 5. Stain the cells with the secondary antibodies for 60 min at room temperature. 6. Briefly stain DNA with 200 μl of 0.2 μg/ml Hoechst 33258 or Hoechst 33342. 7. Wash the cells three times, each time with 200 μl PBT for 60 min. 8. Mount the specimens in 200 μl 50% glycerol in PBS or Aqua Poly-Mount. 9. Examine the immunologically stained samples by phase-contrast and epifluorescent or confocal microscopy.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Culture medium To M3 (BF) medium (see recipe) add 1% (w/v) insulin (see recipe for 100× insulin), 1% (w/v) glutathione (see recipe for 100× glutathione, 10% (v/v) heatinactivated fetal bovine serum (FBS; see recipe), and 10% (v/v) fly extract (see Support Protocol 1). Filter-sterilize the culture medium with a Stericup or syringe filter. Store up to 7 days at 4◦ C. For primary cultures, it is recommended to add a mixture of penicillin/streptomycin (see recipe) to the culture medium.

Glutathione, 100× Dissolve 6.0 g of glutathione (Sigma, cat. no. G6013) in 100 ml of Milli-Q water. Filter sterilize and prepare 1.0-ml aliquots. Store up to 6 months at −20◦ C.

Heat-inactivated FBS Inactivate FBS (ES Cell Qualified; Invitrogen, cat. no. 16141-079) at 60◦ C for 30 min. Somatic Stem Cells

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Insulin, 100× Dissolve 100 mg insulin (Sigma, cat. no. I1882) in 100 ml Milli-Q water, and add 100 μM HCl. Filter sterilize and prepare 1.0-ml aliquots. Store up to 6 months at −20◦ C.

M3 (BF) medium Dissolve Shields and Sang M3 Insect Medium powder (Sigma, cat. no. S8398) in 700 to 800 ml sterilized Milli-Q water. Add 1.0 g potassium glutamate (Sigma, cat. no. G1149) and 0.5 g potassium bicarbonate (Sigma, cat. no. P7682). Adjust to pH 6.85 with 1% NaOH. Add Milli-Q water to make a final volume of 1000 ml. Filter-sterilize the medium with a Stericup (0.22-μm pore size, 1000-ml, Millipore). Store the M3 (BF) medium up to 6 months at 4◦ C.

Microwaved yeast solution Dissolve 30 g of dry yeast (Red Star) in 100 ml of Milli-Q water. Boil the yeast solution once in a microwave oven. Store the microwaved yeast solution up to 2 months at 4◦ C. Boil the microwaved yeast solution every time before use.

Penicillin, 100× Dissolve 6.4 g of penicillin G potassium salt (Sigma, cat. no. P7794) in 100 ml of Milli-Q water. Filter sterilize and prepare 1.0-ml aliquots in 1.5-ml microcentrifuge tubes. Store up to 6 months at −20◦ C.

Streptomycin, 100× Dissolve 1.0 g of streptomycin sulfate salt (Sigma, cat. no. S9137) in 100 ml of Milli-Q water. Filter sterilize and prepare 1.0-ml aliquots in 1.5-ml microcentrifuge tubes. Store up to 6 months at −20◦ C.

COMMENTARY Background Information

Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

Stable cell lines of primordial germ cells (Matsui et al., 1992) and spermatogonial stem cells (Kanatsu-Shinohara et al., 2003) have been successfully cultured in media containing a cocktail of growth factors. However, culture techniques have not been fully developed for the research of primordial germ cells and GSCs in Drosophila melanogaster, which is an excellent model for studying stem cells and their niches. One of the impediments to the use of D. melanogaster is the difficulty of collecting sufficient numbers of purified cells from flies. There are only 30 cells on an average in a wild-type ovary. Another approach for collecting sufficient numbers of purified OSS cells and GSCs is to use flies bearing an ovarian tumorous mutant gene, bam. The wild-type allele of bam functions at the first step of GSC differentiation, and disruption of the bam gene results in tumorous ovaries in which GSCs retain high mitotic activity, and thus continue to expand as the adult female ages. We notice more than 10,000 GSC cells and surprisingly almost the same number of somatic cells in

bam ovarioles of aged females (Niki et al., 2006). The somatic cells are the descendants of two kinds of somatic stem cells: escort stem cells and follicle stem cells, which accumulate in the anterior and middle region, respectively. It should be noted that expanded bam GSCs can differentiate into functional germ cells when they are injected into recipient embryos, and the wild-type allele of the bam gene is artificially expressed during pupal and adult stages (Niki and Mahowald, 2003).

Critical Parameters and Troubleshooting Growth rates of fGS and OSS cells depend on the concentration of FBS and FE. The doubling time is ∼20 hr in a medium with 10% FBS and 10% FE. The cultures should be split 1:2 or 1:4 every week when passaging the cells. Note that cells tend to make cellular clumps and that GSCs are surrounded by somatic cells after confluence (Fig. 2E.1.1E), which sometimes causes the rapid disappearance of GSCs. When cellular clumps start to aggregate, fragment them into pieces by rigorous scraping

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with a 200-μl pipet tip and transfer them into new wells of a culture plate. The GSCs will then begin to expand from the periphery of the fragmented clumps.

Anticipated Results More than 104 GSC and 103 OSS cells can be obtained from one bam female fly. GSCs and OSS cells are associated with each other via DE-cadherin, and signal transduction pathways occur between them (Niki et al., 2006). Thus, it is very useful in the study of stem cell biology to elucidate the mechanisms of maintenance and division of GSCs and the roles of stromal cells at cellular and molecular levels. Because GSCs and OSS cells are of stem cell origin, they are very useful for the biochemical study of the RNAi machinery of Drosophila. Recently, Lau et al. (2009) analyzed the OSS line in detail and found that in addition to miRNAs, it expresses high levels of endo-siRNAs and primary piRNAs. Gene profiling of GSCs is also possible, and thus genes that are expressed in GSCs can be compared between fly and mouse (reviewed by Niki, 2008). OSS cells would also be useful for feeder cells of other germline cells of Drosophila.

Time Considerations The whole process takes several months from isolating GSCs to establishing stable cell lines. Then, cultured cells continue to proliferate dramatically and it takes ∼7 days to reach confluence in each well. It is highly recommended to passage the cells before they reach confluence and not let them overgrow. Otherwise, GSCs disappear when somatic cells surround them. It is easy to expand the cells from a very low density (e.g., <100 cells in a well).

Currie, D.A., Milner, M.J., and Evans, C.W. 1988. The growth and differentiation in vitro leg and wing imaginal disc cells from Drosophila melanogaster. Development 102:805-814. Fuller, M.T. and Spradling, A.C. 2007. Male and female Drosophila germline stem cells: Two versions of immortality. Science 316:402-404. Kanatsu-Shinohara, M., Ogonuki, N., Inoue, K., Miki, H., Ogura, A., Toyokuni, S., and Shinohara, T. 2003. Long-term proliferation in culture and germline transmission of mouse male germline stem cells. Biol. Reprod. 69:612616. Lau, N.C., Robine, N., Martin, R., Chung, W., Niki, Y., Berezikov, E., and Lai, E.C. 2009. Abundant primary piRNAs, endo-siRNAs and microRNAs in a Drosophila ovary cell line. Genome Res. 2009 Jul 14. [Epub ahead of print]. Matsui, Y., Zsebo, K., and Hogan, B.L. 1992. Derivation of pluripotential embryonic stem cells from murine primordial germ cells in culture. Cell 70:841-847. Niki, Y. 2008. In vitro approach of germline stem cells in fly and mouse. In Stem Cell Applications in Disease and Health (W.B. Burnsides and R.H. Ellsley, eds.) pp. 127-149. Nova Science Publishers, New York. Niki, Y. and Mahowald, A.P. 2003. Ovarian cystocytes can repopulate the embryonic germline and produce functional gametes. Proc. Natl. Acad. Sci. U.S.A. 100:14042-14045. Niki, Y., Yamaguchi, F., and Mahowald, A.P. 2006. Establishment of stable cell lines of Drosophila germ-line stem cells. Proc. Natl. Acad. Sci. U.S.A. 103:16325-16330. Robb, J.A. 1969. Maintenance of imaginal discs of Drosophila melanogaster in chemically defined media. J. Cell Biol. 41:876-885. Ueda, R., Ui-Tei, K., Roberts, J., and Cherbas, L. 2007. Standard protocol for establishing cell lines from Drosophila embryos. CGB Technical Report 2007-04. Ui, K., Ueda, R., and Miyake, T. 1987. Cell lines from imaginal discs of Drosophila melanogaster. In Vitro Cell. Dev. Biol. 23:707711.

Literature Cited Cross, D.P. and Sang, J.H. 1978. Cell culture of individual Drosophila embryos. I. Development of wild-type cultures. J. Embryol. Exp. Morphol. 45:161-172.

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Time-Lapse Live Imaging of Stem Cells in Drosophila Testis

UNIT 2E.2

Jun Cheng1 and Alan J. Hunt1 1

University of Michigan, Ann Arbor, Michigan

ABSTRACT This unit describes a protocol for time-lapse live-imaging of stem cells in Drosophila testis. Testis tips are dissected from Drosophila, sliced, and transferred to glass-bottom chambers where the stem cells residing in their native microenvironment can be monitored in real time. This protocol, facilitated with various fluorescence-labeled markers, allows dynamic cellular processes in stem cells to be characterized throughout the cell cycle. C 2009 by John Wiley & Sons, Inc. Curr. Protoc. Stem Cell Biol. 11:2E.2.1-2E.2.8.  Keywords: Drosophila gonad r stem cell r tissue culture r time-lapse live imaging r epifluorescence microscopy

INTRODUCTION The Drosophila melanogaster male gonad has been one of the best model systems to study stem cell biology due to its easily identified anatomy and well-studied signaling pathways. Using this system, many important questions in stem cell biology have been addressed by studying fixed samples. However, the information that can be deduced from fixed samples is limited and potentially ambiguous when reconstructing a dynamic process from static images. This unit describes a protocol of performing time-lapse live imaging of Drosophila testes, which can be used to study the migration patterns of cells or cellular organelles throughout the cell cycle. We focus on the detailed procedures for performing time-lapse live-cell imaging with an inverted epifluorescence microscope, and then describe the method of dissecting and tissue-culturing Drosophila testes in glass-bottom chambers in the Support Protocol. NOTE: This unit assumes that readers have the basic knowledge of laboratory culture of Drosophila; for a more detailed account on Drosophila culture, see Roberts (1998) and Greenspan (2004).

TIME-LAPSE LIVE IMAGING OF DROSOPHILA TESTES This protocol describes the general procedure of performing time-lapse live imaging of Drosophila testes in the glass-bottom culture chamber.

BASIC PROTOCOL

Materials Culture of Drosophila testes tips in petri dishes (Support Protocol) A high-quality inverted microscope with epifluorescence capability (e.g., Zeiss Axiovert 200) 3-axis computer-controlled microscope stage (e.g., Madcity Labs) Automated shutter in the epifluorescence light path (e.g., Uniblitz) Plan-NEOFLUAR 40× objective with NA = 0.75 or AchroPlan 63× objective with NA = 0.8 A highly sensitive CCD camera [e.g., Hamamatsu Electron multiplier (EM) CCD camera] Computer with software for controlling shutter, specimen stage, and image acquisition (e.g., Metamorph or ImageJ) Current Protocols in Stem Cell Biology 2E.2.1-2E.2.8 Published online November 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02e02s11 C 2009 John Wiley & Sons, Inc. Copyright 

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microscope condenser

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Drosophila culture medium cellulose membrane testis tips

glass-bottom petri dish

3-axis movable speciman stage microscope objective

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Figure 2E.2.1 Time-lapse live imaging setup. (A) The petri dish is placed into the chamber holder and assembled. (B) The chamber holder is transferred into the microscope specimen stage. (C) Schematic of the microscope setup, illustrating the relative positions of the sample chamber, the 3-axis specimen stage, the microscope objective, and the microscope condenser. The brightfield image (D) and the epifluorescence image (E) of the Drosophila testis tip of mCherrySas6 flies. Scale bar = 100 μm. (F) Snap shot of EM-CCD setting parameters for mCherry-Sas6 flies.

NOTE: 40× NA = 0.75 and 63× NA = 0.8 objectives provide the necessary depth of field while maintaining sufficient resolution and epifluorescence intensity.

Live Imaging of Drosophila Testis

NOTE: To prevent any photo-bleaching and potential photodamage to the testis tissue, exposure time is minimized. To achieve this goal, a highly sensitive CCD camera and a fast-response shutter in the epifluorescence light pathway are necessary. Moreover, software control of the shutter and CCD camera is required to synchronize the shutter open/close with the timing of the image acquisition.

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1. Place the petri dish with testes (Support Protocol) into the chamber holder (Fig. 2E.2.1A). 2. Place the chamber onto the 3-axis specimen stage (Fig. 2E.2.1B,C). 3. Select the target testis with bright-field microscopy (Fig. 2E.2.1D). 4. Adjust the focus by imaging the target testis with epifluorescence microscopy (Fig. 2E.2.1E). Open and close the shutter promptly to minimize the exposure time when adjusting the focus.

5. Set the proper exposure time and gain for the EM-CCD camera (Fig. 2E.2.1F). To minimize photo bleaching and maintain constant brightness and contrast for fluorescent images over an extended time, the exposure time should be set as short as possible. Although increased camera gain can partially compensate for decreased exposure time, the gain should not be set unnecessarily high, as it increases background noise and EMCCD dark current. For example, the EM gain is set at 500 and exposure time is set at 300 msec for mCherry-Sas6 fly testis (Rusan and Peifer, 2007).

6. Set the interval between exposures. The exposure interval should not be set shorter than necessary; otherwise, photo bleaching may be a problem. The proper exposure interval depends on several factors, at least including the timescale of cellular processes, the stability and brightness of the imaged fluorescent proteins, and the exposure time set in the previous step. For example, the exposure interval is set at 2 min for mCherry-Sas6 fly testis.

7. Start acquiring the image sequence. Multiple image planes can be followed (XYZT sequence) by taking images at different z-focal planes. The movement of the 3-D specimen stage can be controlled using custom-designed software, or integrated with standard imaging packages (Metamorph, ImageJ), and is synchronized with the shutter open/close and image acquisition. Next, the acquired images can be processed and analyzed depending on the experiment. For example, 4-D image sequences (x, y, z, and t) are acquired for mCherry-Sas6 fly testis, and then the centrosome locations (labeled by mCherry-Sas6) are tracked by semi-automatic tracking software (Cheng et al., 2008).

DISSECTING AND TISSUE-CULTURING DROSOPHILA TESTES This protocol describes how to dissect Drosophila testes and how to tissue-culture the testis tips in the glass-bottom petri dish.

SUPPORT PROTOCOL

Materials 70% ethanol Drosophila culture medium (DCM; see recipe) Drosophila male flies Regenerated cellulose membrane (Spectrum Lab) Carbon dioxide flowbed (Genesee Scientific) Stereomicroscope (Leica) Standard dissecting equipment including: Forceps (Dumont #5) Scalpel (Feather #15) Scissors 35-mm glass-bottom Petri dish with 20-mm microwell (MatTek) Parafilm

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Figure 2E.2.2 Drosophila testis tips are prepared for tissue culturing. (A,B) Male Drosophila is dissected and testes are removed in DCM. (C,D) Dissected testes are transferred to new DCM. (E,F) Testis tip (arrowhead) is cut by scalpel. Scale bars = 1 mm.

Prepare for dissection 1. Sterilize all dissecting equipment with 70% ethanol. Sterile dissecting equipment is critical for maintaining a healthy tissue culture condition.

2. Cut regenerated cellulose membrane into 16-mm diameter circles and soak them in DCM at 4◦ C for 24 hr prior to use. The circular regenerated cellulose membranes can be stored in DCM up to 7 days at 4◦ C.

3. Warm DCM to room temperature before use. Prewarming DCM minimizes the temperature shock to Drosophila testes.

Dissect and cut Drosophila testes 4. Anesthetize the fly on a carbon dioxide flowbed. 5. Dissect testis out of the fly abdomen under a stereomicroscope in prewarmed DCM (Fig. 2E.2.2A,B) DCM instead of PBS is used here to minimize the physiological stress on testis tissue during dissection.

6. Transfer the testes into fresh DCM (Fig. 2E.2.2C,D). Live Imaging of Drosophila Testis

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7. Cut off the testis tip with a scalpel (Fig. 2E.2.2E,F) and use the testis tip for experiments. Current Protocols in Stem Cell Biology

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Figure 2E.2.3 Tissue culturing of Drosophila testis tip in the glass-bottom petri dish. (A) Drosophila testis tip is transferred to glass-bottom petri dish in DCM. Inset: a zoom-in view of the testis tip in a drop of DCM. Scale bar = 1 mm. (B) The testis tip is covered by the DCM-presoaked circular regenerated cellulose membrane. (C) DCM is added on the top of the membrane. (D) Additional DCM is added in the petri dish, and the dish is sealed with Parafilm.

Using only the testis tips for experiments eliminates the peristaltic motion associated with the testis tube.

Culture Drosophila testis tip in the glass-bottom chamber 8. Transfer the testis tip into the glass-bottom petri dish (Fig. 2E.2.3A). It may be beneficial that several testis tips are prepared and transferred into one chamber, and then the one with best imaging quality is selected for the experiment.

9. Cover the testis tips with DCM-presoaked circular regenerated cellulose membrane (Fig. 2E.2.3B). The membrane prevents testis tip from floating in the DCM while allowing exchange of dissolved gases and nutrients to sustain the testis tissue.

10. Carefully add 200 to 300 μl DCM to cover the regenerated cellulose membrane (Fig. 2E.2.3C). DCM is added to provide nutrients. By adding DCM on the top of the regenerated cellulose membrane, the membrane is prevented from floating, thus pressing down and immobilizing the testis tissue.

11. Slowly add another 700 to 800 μl DCM inside the periphery of the petri dish, and seal the dish with Parafilm (Fig. 2E.2.3D). Additional DCM inside the petri dish and the seal by Parafilm prevent changes in the concentration of DCM due to evaporation during long-term observation. Testis tips can be kept in the chamber for up to 20 hr in the dark at room temperature before commencing the time-lapse live-imaging experiment.

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Drosophila culture medium (DCM) Schneider’s Drosophila medium (Invitrogen, cat no. 11720-034) 10% (v/v) fetal bovine serum (FBS; Lonza, cat no. 14-501E) 50 U/ml penicillin 50 μg/ml streptomycin Store up to 6 months at 4◦ C COMMENTARY Background Information

Live Imaging of Drosophila Testis

With remarkable prolonged self-renewal ability and the potency to differentiate into specialized cells, stem cells have been the subject of intensive study for their potential medical applications (Morrison et al., 1997; Watt and Hogan, 2000; Morrison and Kimble, 2006). This self-renewal capability also confers on stem cells an intrinsic risk of cancer formation (Groden et al., 1991; Pece et al., 2004; Singh et al., 2004; Clevers, 2005; Clarke and Fuller, 2006). Indeed, a subset of cancer cells has been shown to have stem cell characteristics (Al-Hajj et al., 2003; Singh et al., 2004). On the other hand, excessive differentiation of stem cells is believed to contribute to tissue degeneration and ageing (Van Zant and Liang, 2003; Kirkwood, 2005; Rando, 2006; Brunet and Rando, 2007). Therefore, a better understanding of the regulatory mechanisms balancing self-renewal and differentiation offers the possibility to provide new perspectives on treating cancer and ageing-related diseases. Local microenvironments known as niches govern the fates of the stem cells in many systems (e.g., gonads, hematopoietic system, skin and gut epithelium; Watt and Hogan, 2000; Spradling et al., 2001; Fuchs et al., 2004), and in these tissue architectures, regulatory signals secreted by the niche cells regulate the adjacent stem cells’ self-renewal ability and suppress their differentiation. With well-developed genetic manipulation techniques, Drosophila melanogaster has been one of the most commonly used model organisms in biological research for over a century. Moreover, the stem cell niche in the Drosophila testis is among the best characterized in signaling pathways and anatomy (Brinster, 2002; Lin, 2002; Fuller and Spradling, 2007), offering an ideal model system to study stem cell regulatory mechanisms in the niche microenvironment. Hub cells, a major com-

ponent of the stem cell niche, are located at the apical tip of Drosophila testis and are surrounded by two different types of stem cell populations: germline stem cells (GSCs) and somatic stem cells [i.e., cyst stem cells (CySCs)]. Hub cells specify the stem cell identity of both GSCs and CySCs by secretion of the ligand Unpaired (Upd), which activates the JAK-STAT signaling pathway (Kiger et al., 2001; Tulina and Matunis, 2001; Leatherman and Dinardo, 2008). Each GSC is encapsulated by a pair of CySCs, and this germ-soma important interaction has been shown to play a role in guiding stem cell self-renewal and differentiation (Kiger et al., 2000; Tran et al., 2000; Schulz et al., 2002; Leatherman and Dinardo, 2008). Because of the importance of the interactions among different cell types in regulating functions and fates, the native stem cell niche in Drosophila testis instead of individual cell lines must be cultured and maintained to observe and study normal physiological behavior. Due to the technical challenges of observing stem cells alive in their native niche environment in Drosophila testis, the biomechanical and morphological understanding of stem cell division and mitotic spindle formation had been mostly inferred from examinations of fixed samples. Although these approaches have revealed a great deal, they are limited and potentially misleading due to reliance on reconstructing the dynamics of biomechanical and morphologic events from loosely correlated static images. Following the protocol outlined in this unit, healthy stem cells within the intact niche from Drosophila testis can be maintained for extended time (at least 24 hr), providing a means to study new processes in stem cell biology by time-lapse live imaging. This described protocol was first applied to study the centrosome orientation dynamics in GSCs (Cheng et al., 2008).

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Critical Parameters and Troubleshooting The workstation and all dissecting equipment must be maintained under good sterile conditions to prevent potential contamination of the cultures. Prior to use, the DCM should be warmed up to room temperature to minimize any temperature shock to the tissue. Extra care must be taken when applying DCM onto the regenerated cellulose membrane. Either applying too much DCM or applying DCM too vigorously may float the membrane, and thus the membrane would not serve the function of immobilizing the testis tissue. Exposure time, interval, and total observation time must be coordinated with the characteristics of the targeted fluorescent protein, and every effort must be taken to shorten exposure time and lengthen exposure interval to avoid the potential problems associated with photobleaching and photodamage. For extended periods of observation (e.g., overnight), microscope drift can be a serious challenge. We find drift is most easily suppressed by taking steps to maintain constant temperature in the microscope room.

Anticipated Results This protocol is applicable to study various dynamic processes when relevant molecules are marked by fluorescence proteins. The maintenance of the healthy culture condition over extended time makes it possible to monitor a particular cellular process throughout the cell cycle. Furthermore, we anticipate that this protocol may be modified to obtain timelapse live imaging of other tissue types from Drosophila or other organisms, providing a complimentary means of studying broader biological questions.

Time Considerations To estimate the time necessary for this protocol from dissecting flies to taking time-lapse live imaging, the following four steps need to be taken into account: material preparation, Drosophila dissection and testis preparation, sample chamber preparation, and time-lapse live imaging. In the material preparation, the regenerated cellulose membrane should be cut and soaked in DCM 24 hr before experiment. With experience, it generally takes ∼2 min per fly to perform Drosophila dissection and testis preparation. Five min may be needed to prepare the sample chamber. The duration of time-lapse live imaging is selected in accordance to the experiment.

Acknowledgements We thank Dr. Y. Yamashita for help with the Drosophila studies and Drs. N. Rusan and M. Peifer for generous gifts of fly strain (mCherry-Sas6) shown in Figure 2E.2.1. This study was supported by NIH grant R01GM072006 to A.J.H.

Literature Cited Al-Hajj, M., Wicha, M.S., Benito-Hernandez, A., Morrison, S.J., and Clarke, M.F. 2003. Prospective identification of tumorigenic breast cancer cells. Proc. Natl. Acad. Sci. U.S.A. 100:39833988. Brinster, R.L. 2002. Germline stem cell transplantation and transgenesis. Science 296:21742176. Brunet, A. and Rando, T.A. 2007. Ageing: From stem to stern. Nature 449:288-291. Cheng, J., Turkel, N., Hemati, N., Fuller, M.T., Hunt, A.J., and Yamashita, Y.M. 2008. Centrosome misorientation reduces stem cell division during ageing. Nature 456:599-604. Clarke, M.F. and Fuller, M. 2006. Stem cells and cancer: Two faces of eve. Cell 124:1111-1115. Clevers, H. 2005. Stem cells, asymmetric division and cancer. Nat. Genet. 37:1027-1028. Fuchs, E., Tumbar, T., and Guasch, G. 2004. Socializing with the neighbors: stem cells and their niche. Cell 116:769-778. Fuller, M.T. and Spradling, A.C. 2007. Male and female Drosophila germline stem cells: Two versions of immortality. Science 316:402-404. Greenspan, R.J. 2004. Fly Pushing: The Theory and Practice of Drosophila Genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Groden, J., Thliveris, A., Samowitz, W., Carlson, M., Gelbert, L., Albertsen, H., Joslyn, G., Stevens, J., Spirio, L., Robertson, M., Sargeant, L., Krapcho, K., Wolff, E., Burt, R., Hughes, J.P., Warrington, J., McPherson, J., Wasmuth, J., Le Paslier, D., Abderrahim, H., Cohen, D., Leppert, M., and White, R. 1991. Identification and characterization of the familial adenomatous polyposis coli gene. Cell 66:589-600. Kiger, A.A., White-Cooper, H., and Fuller, M.T. 2000. Somatic support cells restrict germline stem cell self-renewal and promote differentiation. Nature 407:750-754. Kiger, A.A., Jones, D.L., Schulz, C., Rogers, M.B., and Fuller, M.T. 2001. Stem cell self-renewal specified by JAK-STAT activation in response to a support cell cue. Science 294:2542-2545. Kirkwood, T.B. 2005. Understanding the odd science of aging. Cell 120:437-447. Leatherman, J.L. and Dinardo, S. 2008. Zfh-1 controls somatic stem cell self-renewal in the Drosophila testis and nonautonomously influences germline stem cell self-renewal. Cell Stem Cell 3:44-54.

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Lin, H. 2002. The stem-cell niche theory: Lessons from flies. Nat. Rev. Genet. 3:931-940. Morrison, S.J. and Kimble, J. 2006. Asymmetric and symmetric stem-cell divisions in development and cancer. Nature 441:1068-1074. Morrison, S.J., Shah, N.M., and Anderson, D.J. 1997. Regulatory mechanisms in stem cell biology. Cell 88:287-298. Pece, S., Serresi, M., Santolini, E., Capra, M., Hulleman, E., Galimberti, V., Zurrida, S., Maisonneuve, P., Viale, G., and Di Fiore, P.P. 2004. Loss of negative regulation by Numb over Notch is relevant to human breast carcinogenesis. J. Cell Biol. 167:215-221. Rando, T.A. 2006. Stem cells, ageing and the quest for immortality. Nature 441:1080-1086. Rusan, N.M. and Peifer, M. 2007. A role for a novel centrosome cycle in asymmetric cell division. J. Cell Biol. 177:13-20. Roberts, D.B. 1998. Drosophila: A Practical Approach. IRL Press at Oxford University Press, New York. Schulz, C., Wood, C.G., Jones, D.L., Tazuke, S.I., and Fuller, M.T. 2002. Signaling from germ cells

mediated by the rhomboid homolog stet organizes encapsulation by somatic support cells. Development 129:4523-4534. Singh, S.K., Hawkins, C., Clarke, I.D., Squire, J.A., Bayani, J., Hide, T., Henkelman, R.M., Cusimano, M.D., and Dirks, P.B. 2004. Identification of human brain tumour initiating cells. Nature 432:396-401. Spradling, A., Drummond-Barbosa, D., and Kai, T. 2001. Stem cells find their niche. Nature 414:98104. Tran, J., Brenner, T.J., and DiNardo, S. 2000. Somatic control over the germline stem cell lineage during Drosophila spermatogenesis. Nature 407:754-757. Tulina, N. and Matunis, E. 2001. Control of stem cell self-renewal in Drosophila spermatogenesis by JAK-STAT signaling. Science 294:25462549. Van Zant, G. and Liang, Y. 2003. The role of stem cells in aging. Exp. Hematol. 31:659-672. Watt, F.M. and Hogan, B.L. 2000. Out of Eden: stem cells and their niches. Science 287:14271430.

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In Situ Hybridization to Identify Gut Stem Cells

UNIT 2F.1

Alex Gregorieff1 and Hans Clevers1 1

Hubrecht Institute, Utrecht, The Netherlands

ABSTRACT In recent years, considerable effort has been directed towards identifying the repertoire of genes speciÞcally expressed in adult stem cells. In this unit, we describe an in situ hybridization protocol adapted for the analysis of gene expression in the intestinal mucosa. This methodology allows researchers to quickly visualize the expression proÞle of putative stem cell markers with a high degree of sensitivity and resolution. Curr. Protoc. C 2010 by John Wiley & Sons, Inc. Stem Cell Biol. 12:2F.1.1-2F.1.11.  Keywords: (ISH) in situ hybridization r digoxigenin RNA probes r formalin-Þxed parafÞn-embedded r intestinal sections

INTRODUCTION Our ability to visualize, at the cellular level, gene products in whole organisms and tissues represents an essential step towards fully understanding the biological function of any given gene. Two of the most widely used techniques to address this issue include immunohistochemistry (IHC) and in situ hybridization (ISH). Although in comparison, IHC provides a clear advantage in that protein rather than mRNA expression can be detected, ISH is often a more versatile and robust method for monitoring gene expression (in part because it does not rely on having an antibody). In this unit, we describe a method for detecting RNA species in formalin-Þxed, parafÞn-embedded intestinal sections using digoxigenin-labeled RNA probes in combination with alkaline phosphatase (AP)–coupled anti-digoxigenin antibodies. Although initially set up for detecting mRNA expression in intestinal sections, this protocol works equally well for other tissue and cell types. When coupled to other techniques such as microarray proÞling that allow researchers to identify putative stem cell markers, the ISH protocol is particularly useful in validating the expression of candidate genes. The ability to detect intestinal stem cells by in situ hybridization relies on the availability of robust and speciÞc markers. Recently, two populations of gut stem cells have been identiÞed. Lgr5+ stem cells are located at the crypt base intermingled between Paneth cells (Barker et al., 2007), while Bmi1+ cells are positioned immediately above the Paneth cell compartment (Sangiorgi and Capecchi, 2008). Both populations have been visualized by in situ hybridization using speciÞc probes for Lgr5 and Bmi1. In addition, transcriptional proÞling of Lgr5+ cells has uncovered numerous other stem cell markers such as Ascl4 and Olfm4 (van der Flier et al., 2009). Other genes expressed in stem cells include cMyc, Musashi, and Prominin, although these genes are also expressed in the transit amplifying population and therefore are not speciÞcally enriched in stem cells per se. This unit contains two protocols. The Basic Protocol describes preparation of the intestinal tissue and in situ hybridization and the Support Protocol describes preparation of the digoxigenin-labeled RNA probes. Somatic Stem Cells Current Protocols in Stem Cell Biology 2F.1.1-2F.1.11 Published online January 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02f01s12 C 2010 John Wiley & Sons, Inc. Copyright 

2F.1.1 Supplement 12

BASIC PROTOCOL

IN SITU HYBRIDIZATION TO DETECT STEM CELL GENES Prior to performing the in situ hybridization, intestinal samples must be Þrst Þxed, embedded in parafÞn, cut, and mounted on glass slides following standard histological practice. In this initial stage, the most critical step is to ensure proper Þxation of intestinal samples. Once mounted, sections are ready for pretreatments, which will remove parafÞn wax and open the tissue, allowing for greater accessibility of the probe to the mRNA. The sections are then hybridized to digoxigenin-coupled probes (Support Protocol). Following stringent washes to remove excess probe, hybrids are detected with an alkaline phophatase (AP)–coupled antibody recognizing the digoxigenin molecule linked to the probes. Finally, to reveal the signal, an alkaline phosphatase substrate solution is applied to sections.

Materials Rat or mouse intestine, freshly dissected 10% (v/v) neutral buffered formalin (Þxative) Phosphate-buffered saline (PBS; see recipe) 25%, 50%, 75%, 90%, and 100% (v/v) ethanol Xylenes ParafÞn wax Absolute ethanol, 96% ethanol DEPC-treated H2 O HCl Proteinase K Glycine Paraformaldehyde (see recipe) Acetic anhydride solution (see recipe) 20× SSC, pH 4.5 (see recipe) 20× SSC, pH 7.5 (see recipe) Formamide Hybridization solution (see recipe) Digoxigenin-labeled probe (Support Protocol) Tris/NaCl buffer (see recipe) Blocking solution (see recipe) Anti-digoxigenin AP-conjugated antibody (Roche) NTM buffer (see recipe) NBT/BCIP (Sigma) working solution (see recipe) Permanent mounting medium Microtome Tweezers 10-ml syringe and 25-G needle Small cardboard rings and pins Histological slides (Superfrost Plus slides) Temperature-regulated oven Glass jars (e.g., Coplin jars with lids) Covered slide box Coverslips Light microscope

In Situ Hybridization to Identify Gut Stem Cells

Fix the intestinal samples 1. Following dissection, ßush intestines of their fecal content using 10 ml Þxative (formalin). To do so, hold the intestine with tweezers and apply formalin into the gut tube using a needle (25-G) and syringe (10-ml). To avoid damaging the intestinal

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lining, do not apply too much pressure with the syringe. This can be achieved by cutting the intestine into smaller pieces. The intestine deteriorates rapidly after dissection. Thus, it is of the utmost importance to place tissue samples as quickly as possible in Þxative.

2. If Þxing intact intestines, pin these onto a small cardboard ring in spirals (Swiss rolls) and place in at least 25 ml of formalin. Alternatively, to allow for more efÞcient penetration of the Þxative, cut intestine into 2- to 3-cm pieces and then place in ∼25 ml of formalin per whole intestine. 3. Let Þxation in formalin proceed for ∼16 hr with gentle agitation at room temperature (∼22◦ C). 4. Remove Þxative and wash intestines twice, each time in >50 ml PBS.

Dehydrate and parafÞn-embed samples 5. Proceed with standard dehydration and parafÞn embedding protocol (volume for each wash should be >50 ml per whole intestine): 25% ethanol, 15 min at room temperature 50% ethanol, 15 min at room temperature 75% ethanol, 15 min (or overnight at 4◦ C for large pieces) 90% ethanol, 30 min at room temperature 100% ethanol, three times, 1 hr each at room temperature. 6. Replace the last ethanol wash with xylene (>50 ml) and incubate 1 hr at room temperature. Repeat xylene treatments twice, for a total of three times. 7. Place the samples in parafÞn wax (>100 ml). Replace once and incubate overnight in a 55◦ C oven. 8. Replace the parafÞn once and allow samples to solidify at room temperature. 9. Prepare 8- to 10-μM sections and mount on adhesive glass slides. 10. Dry the slides overnight in a 45◦ C oven.

Pretreat sections prior to in situ hybridization 11. Clean glass jars (including covers) suitable for holding glass slides (e.g., Coplin jars with lids) and bake overnight at 200◦ C. 12. Proceed with standard dewaxing and rehydration protocol, by placing slides in jars containing the following solutions for the indicated times:

Xylene, three times, 5 min each 100% ethanol, two times, 5 min each 96% ethanol, one time, 5 min 70% ethanol, one time, 5 min 50% ethanol, one time, 5 min 25% ethanol, one time, 5 min. For each step listed above, it is important to add enough solution in the jar to cover intestinal sections. The exact volumes will depend on the size and shape of jars and the number of slides. Do not let sections dry out during any step.

13. Rinse the slides twice in DEPC-treated H2 O. 14. Treat the slides with 0.2 N HCl for 15 min at room temperature.

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Table 2F.1.1 Incubation Times, Concentration, and Temperature of Proteinase K Treatment for Various Types of Intestinal Sections

Tissue type

Proteinase K concentration in PBS

Time

Temperature

Embryo, E8.5

15 μg/ml

5 min

Room temperature

Embryo, E10.5

15 μg/ml

10 min

Room temperature

Fetal gut, E14.5-E18.5

30 μg/ml

10 min

Room temperature

Adult intestine and colon

30 μg/ml

20 min

37◦ C

15. Incubate the sections with proteinase K in PBS buffer (see Table 2F.1.1 for concentration, temperature, and durations). Place jar with slides in a water bath if required. Optimal concentration and duration of proteinase K treatment should be tested carefully.

16. Rinse the slides in freshly prepared 0.2% glycine/PBS solution for 1 min. Glycine should be added to PBS solution at last moment. Do not store solution for long periods of time.

17. Rinse the slides twice, each time for 1 min in PBS. 18. Post-Þx for 10 min with 4% paraformaldehyde in PBS at room temperature. 19. Rinse the slides three times, each time for 1 min in PBS. 20. Prepare fresh acetic anhydride solution. Shake vigorously and add to the slides in a glass jar. Incubate for 5 min at room temperature. 21. Repeat acetic anyhydride treatment once. 22. Rinse Þve times, each time in PBS for 1 min at room temperature. 23. Rinse two times in 5× SSC, pH 7.5, each time for 1 min at room temperature.

Prehybridize the sections 24. Remove excess solution from slides with tissue and place them in a covered slide box humidiÞed with 5× SSC, pH 7.5/50% formamide. 25. Add enough hybridization solution (∼500 μl) to completely cover the sections. It is not necessary to place a coverslip over section.

26. Incubate the slide box in a 65◦ C oven for at least 1 hr.

Hybridize the sections 27. Remove excess hybridization solution and replace with 300 to 400 μl/slide of hybridization solution containing 500 ng/ml of digoxigenin-labeled probe. Take care that the slides are horizontal in the humidiÞed chamber. Again, no coverslips are necessary.

28. Incubate the slides in an oven at 62◦ to 70◦ C for 24 to 72 hr. Optimal incubation times and temperatures should be empirically tested, although 48 hr at 65◦ C gives satisfactory results for most probes.

In Situ Hybridization to Identify Gut Stem Cells

Wash post-hybridization 29. Remove excess hybridization solution and place in glass jar. Rinse in 2× SSC, pH 7.5, for 1 min at room temperature.

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30. Wash the slides three times for 20 min each at 60◦ to 65◦ C in 50% formamide/2× SSC, pH 4.5. Optimal washing temperature should be tested.

31. Rinse the slides Þve times, each time in Tris/NaCl buffer for 1 min at room temperature.

Detect signal immunologically 32. Remove excess solution from the slides with tissue and place them in a covered slide box humidiÞed with water. 33. Apply blocking solution over sections (500 μl per slide) and incubate at room temperature for at least 30 min. 34. Dilute sheep anti-digoxigenin antibody 1/2000 in blocking solution. 35. Remove blocking solution and replace with antibody solution (400 μl per slide). Incubate overnight or longer at 4◦ C. 36. Wash the slides Þve to seven times, each time in Tris/NaCl buffer for 1 min at room temperature. 37. Wash the slides two to three times, each time in NTM buffer for 1 min at room temperature. 38. Add NBT/BCIP working solution to sections in a humidiÞed slide box. Incubate for up to 24 hr at room temperature, keeping the slides in the dark. 39. Wash the slides twice, each time in PBS for 1 min at room temperature. 40. Dehydrate the sections as follows:

Rinse sections in H2 O Rinse in 70% (v/v) ethanol Rinse in 90% (v/v) ethanol Rinse twice in 100% (v/v) ethanol Rinse twice in xylenes. Proceed quickly through each step. Part of the signal may be lost with extensive washes in ethanol. To avoid signal loss, sections may be visualized under the microscope prior to dehydration. In this case, a few drops of glycerol are added to the slide followed by a coverslip. This method will also reduce the diffusion of the signal resulting from dehydration in ethanol.

41. Apply mounting medium and place a coverslip over the section. 42. Examine the slides with a light microscope. See Anticipated Results for a description of a positive signal.

GENERATION OF DIGOXIGENIN RNA PROBES The generation of digoxigenin RNA probes is achieved by an in vitro transcription reaction of linearized template DNA using T7, T3, or SP6 RNA polymerases. During the in vitro transcription reaction, digoxigenin-coupled UTPs are incorporated into the RNA probe. To generate anti-sense probes that will recognize sense mRNA, template DNA is cut using a restriction enzyme that creates a 5 overhang (avoid 3 overhangs) at the 5 end of the cDNA. Ensure that following the digestion the T7, T3, or SP6 promoter is at the 3 end of the template DNA. Accordingly, to generate sense probes, template DNA can be cut at the 3 end.

SUPPORT PROTOCOL

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A useful source of probes for in situ hybridization can be found through the IMAGE consortium (http://image.hudsonalpha.org/). Using Web-based tools speciÞc cDNAs can be searched and ordered. For detecting stem cells, we recommend generating a full-length probe of Olfm4. When compared to other stem cell markers (e.g., Lgr5, Ascl4, or Bmi1), Olfm4 gives strong and highly reproducible signals, and for this reason, it is perhaps the most useful marker to detect Lgr5+ stem cells by ISH.

Materials Plasmid for gene of interest (e.g., see Table 2F.1.3) Restriction endonuclease and buffer (Bloch and Grossman, 1995) Agarose 3 M sodium acetate, pH 5.2 Phenol/choloroform Absolute ethanol 70% (v/v) ethanol 10 × transcription buffer (Roche) Dithiothreitol (DTT) 10 × Dig RNA labeling mix (Roche) RNase inhibitor (Fermentas) T7 or T3 or SP6 RNA polymerases (Roche) DEPC-treated H2 O Rnase-free DNaseI (Fermentas), optional 4 M LiCl Formamide 1.5-ml microcentrifuge tubes RNA puriÞcation columns (RNeasy Mini Kit, Qiagen) Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2000) and digestion of DNA with restriction enzymes (Bloch and Grossman, 1995) Digest plasmid DNA 1. Digest 10 μg of plasmid DNA with the appropriate restriction enzyme (Bloch and Grossmann, 1995). 2. Ensure that template DNA is completely linearized by running an aliquot on 1% agarose gel (Voytas, 2000). 3. Add 1/10 volume of 3 M sodium acetate, pH 5.2, to the digest and extract once with 50:50 (v/v) phenol/chloroform mixture and once again with chloroform to remove trace amounts of phenol. 4. Precipitate DNA by adding 2.5 vol of absolute ethanol and centrifuge 10 min at 14,000 rpm, 4◦ C, in a benchtop microcentrifuge. 5. Wash the pellet in 500 μl of 70% ethanol and resuspend in 15 μl water. To avoid phenol/chloroform extraction and ethanol precipitation steps, linearized DNA can be puriÞed using commercially available columns.

Perform in vitro translation with digoxigenin dUTP 6. Prepare in vitro transcription reaction in a 1.5-ml microcentrifuge tube (20-μl reaction volume) as follows: In Situ Hybridization to Identify Gut Stem Cells

1 to 2 μg of linearized DNA (from step 5) 1× transcription buffer (usually supplied by manufacturer of RNA polymerases) 2 μl of 0.1 M DTT 2 μl 10× Dig RNA labeling mix

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20 U of RNase inhibitor 30 U of T7 or T3 or SP6 RNA polymerase. Use the polymerase that corresponds to the promoter at the 3 end of the linearized DNA template. See the Introduction above.

7. Incubate the reaction at 37◦ C for 3 hr. 8. Clean-up cRNA products by using commercially available RNA puriÞcation columns. Elute samples from columns with 50 μl DEPC-treated H2 O. As an alternative to using RNA puriÞcation columns researchers may follow these steps to clean up cRNA reaction. a. Add 1 U RNase-free DNaseI to the in vitro transcription reaction, and incubate samples 15 min at 37◦ C. b. Add 2.5μl 4 M LiCl and 75 μl ethanol to the reaction; store at −70◦ C for 20 min. c. Microcentrifuge 5 min at 14,000 rpm, 4◦ C. Wash the pellet in 500 μl of 70% ethanol and resuspend in 50 μl DEPC-treated H2 O.

9. Remove 1 μl of the puriÞed probe to measure the concentration and 3 μl for electrophoresis on standard 1% agarose/ethidium bromide gel (Voytas, 2000). Add an equal volume of 100% formamide to the remaining probe and store at −70◦ C.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Acetic anhydride solution 0.25% (v/v) acetic anhydride in 0.1 M triethanolamine, pH 8.0 Add acetic anhydride solution immediately before use. Do not store! 0.1 M triethanolamine, pH 8.0, may be stored at room temperature

Blocking solution 1% (w/v) blocking powder (Roche) in 1× Tris/NaCl buffer (see recipe). Heat at 65◦ C to dissolve. Usually prepared fresh, but can be stored up to 3 months at −20◦ C.

Hybridization solution 50% (v/v) formamide 5× SSC, pH 4.5 2% (w/v) blocking powder (Roche) 0.05% (w/v) CHAPS 5 mM EDTA 50 μg/ml heparin 1 μg/ml yeast RNA Heat at 65◦ C to dissolve Usually prepared fresh, but can be stored up to 3 months at −20◦ C.

NBT/BCIP solution NBT stock solution: 10 mg of NBT (Sigma) in 1 ml of H2 O continued

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Heat and vortex if required Protect from light and store up to 6 months at −20◦ C BCIP stock solution: 25 mg of BCIP (Sigma) in 500 μl DMF (dimethylformamide) Store up to 6 months at −20◦ C Working solution: 10 ml NTM buffer (see recipe) 25 μl 1 M levamisole 333 μl NBT stock solution 35 μl BCIP stock solution Prepare fresh NTM buffer 0.1 M Tris·Cl, pH 9.5 0.1 M NaCl 0.05 M MgCl2 Prepare fresh Paraformaldehyde (PFA) in PBS, 4% (w/v) Heat at 65◦ C to dissolve Prepare fresh Phosphate-buffered saline (PBS) For 1 liter: 8 g NaCl 0.2 g KCl 2.68 g Na2 HPO4 ·7H2 O 0.24 g KH2 PO4 800 ml H2 O Adjust pH to 7.4 with HCl Adjust volume to 1 liter with H2 O Store up to 6 months at room temperature SSC, 20× For 1 liter: 175.3 g NaCl 88.2 g sodium citrate·2H2 O 800 ml H2 O Adjust pH to 7.5 or 4.5 with HCl Adjust volume to 1 liter with H2 O Store up to 6 months at room temperature When SSC is combined with formamide, pH 4.5 is used. This will ensure that the formamide/ SSC solution will be pH neutral.

Tris/NaCl buffer

In Situ Hybridization to Identify Gut Stem Cells

0.1 M Tris·Cl, pH 7.5 0.15 M NaCl 0.1 % (v/v) Tween 20 Store up to 6 months at room temperature

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COMMENTARY Background Information More than 30 years ago, Cheng and Leblond proposed that intestinal stem cells are located at the base of the crypts of Lieberk¨uhn (Cheng and Leblonde, 1974a,b). These socalled crypt base columnar (CBC) cells were found to be highly proliferative cells, capable of giving rise to the four differentiated cell types of the intestinal epithelium (i.e., enterocytes, goblet cells, Paneth cells, and enteroendocrine). Around the same time, however, pulse-chase experiments with tritiated thymidine led Potten and colleagues to different conclusions. Indeed these researchers demonstrated that label-retaining cells were preferentially located immediately above the Paneth cell compartment (Potten et al., 1974, 2002; Potten, 1977). This population of stem cells is referred to as +4 cells given their position along the crypt-villus axis relative to the bottom of the crypts. Although several question marks still persist regarding the properties of these two populations (+4 cells versus CBC cells), recent knock-in strategies have led investigators in the Þeld to more reliably identify and track intestinal stem cells. In the Þrst model, Barker et al. labeled CBC cells using mice engineered to carry a GFP and a tamoxifen inducible Cre in the Lgr5 locus (Barker et al., 2007). In follow up studies, the Lgr5-GFP-CreER allele was utilized to identify novel stem cell markers based on transcriptional proÞling of sorted GFP-expressing CBC cells (van der Flier et al., 2009). A similar approach allowed Sangiorgi and Capechi to speciÞcally label +4 cells by introducing the CreER enzyme in the locus of the Polycomb family member, Bmi1 (Sangiorgi and Capecchi, 2008). In either case, crossing these lines with the R26RLacZ reporter mice to permanently mark +4 cells or CBC cells demonstrated unequivocally that both cell populations are long-lived and generate all four cell types of intestinal epithelium.

Critical Parameters and Troubleshooting The optimal length of the probe should be determined empirically. In most cases, fulllength probes give satisfactory results. Note however that the yield and quality of the in vitro transcription reaction may be limited when using very large template DNA (>3 kB). PCR fragments with the appropriate promoter added to the 3 end can also be used as an alter-

native to plasmid DNA for generating template DNA. Although alkaline phosphatase (AP)–based detection schemes offer high sensitivity, one caveat in the intestine is the fact that the epithelium harbors a high level of endogenous AP activity. To circumvent this problem it is important to inactivate endogenous AP activity by pretreating sections in 0.2 N HCl. The drug levamisole is an effective way to reduce endogenous AP activity when performing in situ hybridization on whole embryos. In the case of adult intestinal material, however, levamisole is not sufÞcient to block AP activity. Incomplete inactivation of enodogenous AP appears as a thin apical staining of enterocytes. It is also believed that HCl treatments may favor accessibility of the probe to target sequences by extracting mRNA-bound proteins. Another commonly used treatment is acetylation, using acetic anhydride (0.25%) in triethanolamine. This treatment is also thought to be important for decreasing background, but it also appears to inactivate RNases and may help in producing a stronger signal. For troubleshooting information for in situ hybridization, see Table 2F.1.2.

Anticipated Results For probe synthesis, cRNA samples run on normal agarose gels usually give a prominent but diffused band. Occasionally, certain probes may give more of a smear. One can expect to obtain a yield of 7 to 15 μg of cRNA. Following ISH, the intensity and speciÞcity of the signal will depend on the expression pattern and levels of the gene of interest. Generally, for the intestine, genes that are expressed in differentiated epithelial cell types (enterocytes, goblet cells, enteroendocrine cells, and Paneth cells) are readily detected within a few hours of development. Genes exclusively expressed in progenitor/stem cells may require longer development times and signal may be weaker. Generally, the staining pattern appears as a uniform blue or purple deposit within the contours of a given cell. Stem cell markers should result in discrete staining of CBC cells and/or +4 cells and no staining should be observed in the adjacent Paneth cells. The latter can be easily distinguished from CBC or +4 cells by their large size and granular morphology. Occasionally, staining will appear as discrete dots in or around the Paneth cells. This type of pattern should be considered as background.

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Table 2F.1.2 Troubleshooting Guide to In Situ Hybridization to Identify Intestinal Stem Cells

Problem

Possible cause

Solution

Poor histology

Fixation inadequate

Make sure intestines are immediately Þxed following dissection. Take care when ßushing intestines or ßush directly with formalin instead of PBS. Reduce time and concentration of proteinase K treatment

Proteinase K treatment too long Very strong and spotty staining in Paneth, goblet, or enteroedocrine cells

UnspeciÞc binding to secretory products of Paneth, goblet, or enteroendocrine cells

Prepare new intestinal sections. Use a different probe.

Weak or no staining

Temperature of hybridization or post-hybridization washes is too stringent

Prepare new intestinal sections. Lower hybridization/washing temperature. Use a different probe.

High background

Temperature of hybridization or post-hybridization washes is not stringent enough

Raise hybridization/washing temperature. Use a different probe.

Table 2F.1.3 Useful Epithelial, Cell-Type SpeciÞc Markers for In Situ Hybridization in the Small Intestine

Cell type

Gene

Enterocytes

Fabp1, Fabp2

Goblet cells

Gob5, Tff3

Enteroendocrine cells

Chromogranin B

Paneth cells

Cryptdins

Entire epithelium

Villin, Tcf4

Proliferative crypt epithelial cells

c-Myc, c-Myb

Early progenitors/stem cells

Axin2, Lgr5, Olfm4

Table 2F.1.3 lists genes from which suitable probes can be generated to mark certain cell types of the intestine.

Time Considerations The generation of labeled probe including digestion and in vitro transcription reactions can be completed in a single day or a day and a half. The Þxation, dehydration, and parafÞn embedding of intestinal samples require at least 24 to 36 hr. The in situ hybridization protocol takes a minimum of 4 days depending on the length of hybridization and duration of staining procedure (see individual steps).

Literature Cited In Situ Hybridization to Identify Gut Stem Cells

Barker, N., van Es, J.H., Kuipers, J., Kujala, P., van den Born, M., Cozijnsen, M., Haegebarth, A., Korving, J., Begthel, H., Peters, P.J., and Clevers, H. 2007. IdentiÞcation of stem cells in

small intestine and colon by marker gene lgr5. Nature 449:1003-1007. Bloch, K.D. and Grossmann, B. 1995. Digestion of DNA with restriction endonucleases. Curr. Protoc. Mol. Biol. 31:3.1.1-3.1.21. Cheng, H. and Leblond, C.P. 1974a. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. I. Columnar cell. Am. J. Anat. 141:461-479. Cheng, H. and Leblond, C.P. 1974b. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. V. Unitarian theory of the origin of the four epithelial cell types. Am. J. Anat. 141:537561. Potten, C.S. 1977. Extreme sensitivity of some intestinal crypt cells to x and gamma irradiation. Nature 269:518-521. Potten, C.S., Kovacs, L., and Hamilton, E. 1974. Continuous labelling studies on mouse skin and intestine. Cell Tissue Kinet. 7:271-283.

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Potten, C.S., Owen, G., and Booth, D. 2002. Intestinal stem cells protect their genome by selective segregation of template DNA strands. J. Cell Sci. 115:2381-2388. Sangiorgi, E. and Capecchi, M.R. 2008. Bmi1 is expressed in vivo in intestinal stem cells. Nat. Genet. 40:915-920. van der Flier, L.G., van Gijn, M.E., Hatzis, P., Kujala, P., Haegebarth, A., Stange, D.E., Begthel, H., van den Born, M., Guryev, V., Oving, I., van Es, J.H., Barker, N., Peters, P.J., van der Wetering, M, and Clevers, H. 2009. Transcription factor achaete scute-like 2 controls intestinal stem cell fate. Cell 136:903-912. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9.

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Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

UNIT 2G.1

Ivan Bertoncello1 and Jonathan McQualter1 1

Lung Regeneration Laboratory, The Department of Pharmacology, University of Melbourne, Victoria, Australia

ABSTRACT Adult mouse lung epithelial stem/progenitor cells (EpiSPC) can be defined in vitro as epithelial colony-forming units that are capable of self-renewal, and which when cocultured with lung mesenchymal stromal cells (MSC) are able to give rise to differentiated progeny comprising mature lung epithelial cells. This unit describes a protocol for the prospective isolation and in vitro propagation and differentiation of adult mouse lung EpiSPC. The strategy used for selection of EpiSPC and MSC from adult mouse lung by enzymatic digestion and flow cytometry is based on the differential expression of CD45, CD31, Sca-1, EpCAM, and CD24. The culture conditions required for the differentiation (co-culture with MSC) and expansion (stromal-free culture with FGF-10 and HGF) of C 2011 by John EpiSPC are described. Curr. Protoc. Stem Cell Biol. 16:2G.1.1-2G.1.12.  Wiley & Sons, Inc. Keywords: lung epithelium r stem cells r colony-forming assay

INTRODUCTION This unit describes isolation and culture of adult mouse lung epithelial stem/progenitor cells (EpiSPC). EpiSPC are defined as colony-forming units (CFU) that have the capacity for self-renewal and are able to generate progeny of differentiated lung epithelial cells. Basic Protocol 1 describes a cell-fractionation method using flow cytometry for isolating EpiSPC and mesenchymal stromal cells (MSC) from enzymatically digested adult mouse lung tissue, based on their differential expression of CD45, CD31, Sca-1, EpCAM, and CD24 (McQualter et al., 2010). Basic Protocols 2 and 3 describe culture methods for the differentiation, maintenance, and propagation of adult lung EpiSPC, respectively. When co-cultured with MSC in an organotypic three-dimensional Matrigel culture, EpiSPC generate colonies comprising mature differentiated lung epithelial cells. In stromal-free cultures, EpiSPC undergo clonal proliferation when supplemented with FGF-10 and hepatocyte growth factor (HGF), and can be enzymatically dissociated and passaged using this culture setup, demonstrating their high proliferative capacity and ability to self-renew.

ISOLATION OF EPITHELIAL STEM/PROGENITOR CELLS AND MESENCHYMAL STROMAL CELLS FROM ADULT MOUSE LUNG

BASIC PROTOCOL 1

This protocol is used for the dissociation of adult mouse lung tissue and the enrichment of cell fractions containing EpiSPC and MSC. A combination of mechanical and enzymatic dissociation is used to prepare a single-cell suspension of lung cells, followed by a pre-enrichment step using discontinuous density gradient centrifugation to remove high-density cells such as erythrocytes, mature myeloid cells, and cellular debris. Flow cytometry is then used to identify and sort subpopulations of cells enriched for either EpiSPC or MSC, based on the expression of defined cell-surface markers (McQualter et al., 2009, 2010). Somatic Stem Cells Current Protocols in Stem Cell Biology 2G.1.1-2G.1.12 Published online January 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc02g01s16 C 2011 John Wiley & Sons, Inc. Copyright 

2G.1.1 Supplement 16

Materials Collagenase, type 1 (Worthington) Phosphate buffered saline (PBS; Sigma, cat. no. P-3813) Adult mice (strain C57BL/6; 6 to 12 weeks; 18 to 22 g; male or female) PBS with 2% (v/v) fetal bovine serum (FBS; JRH Biosciences, batch tested) Nycoprep density gradient medium (1.077 g/cm3 , 265 mOsm (see recipe) Fluorochrome-conjugated antibodies against: EpCAM (clone G8.8) CD24 (clone M1/69) Sca-1 (clone E13-161.7) CD45 (clone 30-F11) CD31 (clone 390) Viability dye (4 ,6-diamidino-2-phenylindole (DAPI), propidium iodide (PI), or FluoroGold (Fluorochrome LLC, http://www.fluorochrome.com) Dissecting equipment 15- and 50-ml conical polypropylene centrifuge tubes (e.g., BD Falcon) 60-mm-diameter Petri dishes Thermomixer (Eppendorf) 18-G and 21-G needles 5-ml and 20-ml syringes 40-μm cell strainer (BD Biosciences) Refrigerated centrifuge (with capacity for 5-ml, 15-ml and 50-ml tubes) Sysmex KX-21N cell counter (Sysmex Corporation; http://www.sysmex.com/) or hemacytometer counting chamber and trypan blue (UNIT 1C.3) Sterile mixing cannulas (Unomedical, cat. no. 500.11.012; http://www.unomedical.com) 5-ml FACS tubes with 35-μm cell strainer caps (BD Biosciences), sterile Flow cytometer Additional reagents and equipment for euthanasia of mice (Donovan and Brown, 2006), determining viable cell concentration (UNIT 1C.3), and flow cytometry (Robinson et al., 2010) Dissociate lung tissue 1. Prepare 1 mg/ml collagenase type I solution in sterile PBS and preheat to 37◦ C, allowing 3 ml per whole mouse lung. 2. Euthanize mouse by cervical dislocation (Donovan and Brown, 2006), open mouse abdominal cavity, and sever major arteries/vessels behind intestines to exsanguinate animal. 3. Open thoracic cavity, carefully dissect out the lungs, place into a 15-ml tube containing 10 ml chilled PBS, and agitate to rinse out excess blood. If preparing multiple lungs, pool lungs and rinse into a 50-ml tube (up to five lungs per tube). This protocol has been optimized for the isolation of distal lung cells. All upper airways are carefully removed at this point.

4. Transfer lungs into a second 60-ml tube of fresh chilled PBS and rinse lungs well by gentle agitation. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

The lungs should become pale when adequately rinsed. Rinse a third time if necessary.

5. Transfer the lungs into a sterile 60-mm-diameter Petri dish and finely mince the lungs using fine dissecting scissors or a single-sided razor blade.

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Prepare single-cell suspension 6. Transfer the minced tissue into a 15-ml conical centrifuge tube and add the preheated collagenase type I solution (3 ml per lung) to the tube. If processing multiple lungs, use a 50-ml centrifuge tube (up to five lungs per tube).

7. Place the tube in a Thermomixer (Eppendorf) and agitate (750 rpm, 37◦ C) for 30 min. 8. Remove the tube from the Thermomixer and triturate with an 18-G needle attached to a 5-ml syringe until chunks of tissue are mostly dissociated. If processing multiple samples, use a 20-ml syringe.

9. Return tube to the Thermomixer and agitate (750 rpm, 37◦ C) for a further 15 to 30 min until most of the lung tissue fragments are digested. When tissue digestion is complete, you should be unable to see chunks of pink lung tissue, although clumps of extracellular matrix will remain visible as white strands in the suspension.

10. Following digestion, remove the tube from the Thermomixer and triturate with a 21-G needle attached to a 20-ml syringe to generate a single-cell suspension. 11. Strain the tissue digest through a 40-μm cell strainer into a clean, sterile 50-ml centrifuge tube. 12. Resuspend the tissue digest in 50 ml of PBS containing 2% fetal bovine serum and centrifuge 5 min at 400 × g, 4◦ C. 13. Remove supernatant and resuspend the cell pellet in 50 ml of PBS containing 2% fetal bovine serum. To maximize cell recovery, supernatant can be collected in a 50-ml tube and recentrifuged.

14. Centrifuge tube(s) 5 min at 400 × g, 4◦ C. 15. Remove supernatant(s) and resuspend the cell pellet(s) in PBS containing 2% fetal bovine serum (5 ml per lung). 16. Count the cells and calculate cell concentration. A Sysmex KX-21N automated cell counter can be used to count cells, or alternatively a hemacytometer can be used with trypan blue (UNIT 1C.3) to exclude nonviable cells. Average cell yield per lung is 2.2 × 107 cells.

Prepare low-density cell fraction 17. Transfer the cell suspension (5 ml) into a 15-ml centrifuge tube and underlay the suspension with 3 ml of Nycoprep (1.077 g/cm3 ) using a sterile mixing cannula attached to a 20-ml syringe. If processing multiple lungs, the equivalent cell suspension from four lungs (20 ml) can be transferred to a 50-ml centrifuge tube and underlayed with 10 ml of Nycoprep.

18. Centrifuge the gradients 15 min at 600 × g, 21◦ C, with the brake off. Gradient centrifugation will result in most high-density cells (i.e., erythrocytes, mature myeloid cells, and cellular debris) passing through the Nycoprep, leaving an enriched band of low-density cells at the Nycoprep interface.

19. Harvest low-density cells from the interface between the PBS layer and the Nycoprep solution using a 10-ml pipet and transfer into a 50-ml centrifuge tube (Fig. 2G.1.1A). If processing multiple lungs, the low-density cell fractions can be pooled at this point. Average low-density cell yield per lung is 6.9 × 106 cells.

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Figure 2G.1.1 Cell fractionation strategy used to enrich and sort EpiSPC and MSC from enzymatically digested lung tissue. (A) Photographic image of low-density cell band from density gradient centrifugation. (B) Gating strategy inclusion of PIneg viable cells, and (C) exclusion of CD45neg CD31neg cells. (D) Gating strategy for sorting CD45neg CD31neg EpCAMneg Sca-1hi MSC. (E) Gating strategy for sorting CD45neg CD31neg EpCAMhi CD24low EpiSPC.

20. Top up tube(s) containing the low-density cells to a total volume of 50 ml with PBS containing 2% fetal bovine serum and centrifuge 5 min at 400 × g, 4◦ C. 21. Discard supernatant(s) and resuspend the cell pellet(s) in 50 ml PBS containing 2% fetal bovine serum. 22. Centrifuge the tube(s) 5 min at 400 × g, 4◦ C. 23. Discard supernatant(s) and resuspend the cell pellet(s) in 5 ml per lung of PBS containing 2% fetal bovine serum. 24. Count the cells and calculate cell concentration (UNIT 1C.3).

Prepare cells for flow cytometry 25. Aliquot 100,000 to 200,000 cells into a 5-ml FACS tube for each of the compensation tubes and 500,000 cells for isotype control tubes. Set aside remaining cells in a 15-ml conical centrifuge tube (cells for sorting). Compensation tubes should be prepared for each of the antibodies and fluorochromes to be used in the sort strategy. An aliquot of unstained cells should also be prepared.

26. Wash all samples, including cells for sorting, with PBS containing 2% fetal bovine serum, centrifuge 5 min at 400 × g, 4◦ C, and discard supernatant. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

27. Resuspend cell pellets in compensation tubes in 50 μl of each optimally pretitered antibody, and resuspend cells for sorting at 5 × 106 cells per 100 μl of optimally pretitered antibody combination. Use PBS containing 2% fetal bovine serum to dilute antibodies. If streptavidin-conjugated antibodies are used, PBS containing 0.5% (w/v) BSA should be used as a diluent to prevent reactivity of streptavidin with serum biotin.

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28. Incubate all samples on ice in the dark for 20 min. 29. Wash cells in PBS containing 2% fetal bovine serum, centrifuge 5 min at 400 × g, 4◦ C, and discard supernatant. Repeat steps 26 to 29 if secondary or tertiary antibody labeling steps are required.

30. Resuspend cells in compensation tubes in 300 μl and cells for sorting and isotype control tubes at 10–15 × 106 cells/ml in PBS containing 2% fetal bovine serum and an appropriate viability dye. Depending on the flow cytometer setup and laser configuration, DAPI (0.5 μg/ml), PI (1 μg/ml), or Fluorogold (2 μM) may be used as a viability dye. Viability dyes should be included in all compensation, control, and sort tubes.

31. To remove any cell clumps which may block the flow cytometer, aliquot samples into 5-ml FACS tubes with cell strainer caps (35-μm, sterile). To filter larger volumes of cells for sorting, a 40-μm nylon-mesh cell strainer can be used.

Set up for FACS Detailed protocols for flow cytometry are provided in Robinson et al. (2010). 32. Prepare collection tubes containing PBS with 2% fetal bovine serum. Cells can be collected into microcentrifuge tubes (containing 200 μl of PBS/2% fetal bovine serum) or 5 ml FACS tubes (containing 1 ml of the PBS/serum).

33. Set up flow cytometer with a large (90 to 100-μm) nozzle and stabilize the flow stream under low pressure (30 to 40 psi). 34. Set the compensation settings using single-color control tubes. When setting the compensation using single-color control tubes containing lung cells stained with the appropriate antibodies and fluorochromes, it is important to take into consideration the autofluorescence of lung cells and avoid overcompensation (see Critical Parameters).

35. To sort lung EpiSPC and MSC, set sequential gates for selection of viable cells (Fig. 2G.1.1B) followed by exclusion of hematopoietic (CD45) and endothelial (CD31) cells (Fig. 2G.1.1C), prior to selection of EpCAMhi CD24low EpiSPC (Fig. 2G.1.1D) and/or EpCAMneg Sca-1hi MSC (Fig. 2G.1.1E). Isotype controls should be used to account for non-specific antibody staining in setting gates to identify and isolate antibody-positive cells.

36. Sort cells into collection tubes containing PBS containing 2% fetal bovine serum.

ORGANOTYPIC CULTURE OF LUNG EPITHELIAL STEM/PROGENITOR CELLS (DIFFERENTIATION CULTURE)

BASIC PROTOCOL 2

This protocol is used for the identification and characterization of EpiSPC. During culture, epithelial CFU (Epi-CFU) proliferate and differentiate to form complex lineage-restricted or multipotent epithelial colonies comprising alveolar cells, airway cells, or cells of mixed lineage. The growth of these colonies requires seeding EpiSPC in a three-dimensional extracellular matrix (Matrigel) in co-culture with lung-derived MSC (Fig. 2G.1.2A,B). These cultures can be re-fed and maintained over 2 weeks, after which cells begin to die and colonies deteriorate. This culture technique has been developed for EpiSPC sorted on the phenotype CD45neg CD31neg EpCAMhi CD24low , but can also be used to identify EpiSPC from heterogeneous cell fractions. MSC are sorted on the phenotype of CD45neg CD31neg EpCAMneg Sca-1hi .

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Figure 2G.1.2 (A) Schematic representation of organotypic culture setup for EpiSPC differentiation. (B) Image of 24-well tissue culture plate with culture inserts. (C) Whole-well image of colonies after 2 weeks in culture. (D) Bright-field and (E) dark-field images depicting representative colony subtypes, including (i) airway, (ii) alveolar, and (iii) mixed colonies.

NOTE: All procedures are performed in a sterile class II biological hazard flow hood or a laminar-flow hood. All solutions, reagents, media and equipment used to process and culture EpiSPC must be sterile, and proper aseptic technique should be used.

Materials Epi-CFU medium (see recipe) Matrigel (standard concentration; BD Biosciences) Adult lung EpiSPC (CD45neg CD31neg EpCAMhi CD24low ) cells (Basic Protocol 1) Adult lung MSC (CD45neg CD31neg EpCAMneg Sca-1hi ) cells (Basic Protocol 1) Refrigerated centrifuge Millicell-CM culture inserts (0.4-μm membrane, 12-mm diameter, hydrophilic PTFE, Millipore) 24-well flat-bottom tissue culture plates Triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ; see Critical Parameters), humidified Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

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Additional reagents and equipment for determining viable cell concentration (UNIT 1C.3) Prepare organotypic cultures of lung EpiSPC 1. Following flow cytometric cell sorting, wash the sorted EpiSPC and MSC separately in 5 ml/tube Epi-CFU medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. Current Protocols in Stem Cell Biology

2. Resuspend individual cell pellets in Epi-CFU medium (1 to 2 ml per lung) and take an aliquot to determine cell concentration (UNIT 1C.3). Cell counts can be performed using an automated cell counter or a hemacytometer (UNIT 1C.3).

3. After determining cell concentrations, take aliquots of EpiSPC and MSC and combine so that the final mixed cell suspension contains the desired number of cells for culture. Allow for 100 μl per well. For optimal Epi-CFU growth supporting capacity, MSC (CD45neg CD31neg EpCAMneg Sca1hi ) should be used at 2 × 106 cells/ml. The concentration of sorted cells seeded for detection of EpiSPC depends on their level of enrichment in the sorted fraction. Ideally, cells should be seeded at a concentration which will generate about 20 colonies per Millicell insert (∼500 cells). The colony-forming efficiency of CD45neg CD31neg EpCAMhi CD24low EpiSPC from adult (8- to 12-week old) C57Bl/6 mice is typically 1 in 23.

4. Centrifuge EpiSPC/MSC mix 5 min at 400 × g, 4◦ C, and aspirate medium.

Prepare Matrigel suspension cultures 5. Resuspend cell pellet in Matrigel diluted 1:1 with Epi-CFU medium. Allow 100 μl per well. Ensure matrix always stays on ice; otherwise, it will solidify.

6. Gently mix the Matrigel cell suspension. It is important to avoid creating bubbles. This can be achieved by holding the tube in the center of a Vortex mixer to create a swirling motion.

7. Place 12-mm Millicell-CM inserts in 24-well culture plates. 8. Add 90 μl of Matrigel cell suspension on top of the filter membrane of a Millicell-CM insert and place plate in incubator (37◦ C) for 5 min to allow matrix to set. Be careful to avoid creating bubbles in the suspension.

9. Remove plate from incubator and add 400 μl of Epi-CFU medium per well around the insert. 400 μl is just enough to allow the medium to touch the bottom of the insert, allowing diffusion of medium into the Matrigel without submerging the Matrigel culture. This semi-dry state of the Matrigel is essential for epithelial colony-formation.

10. Incubate cell cultures in a humidified 37◦ C triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ) and change to fresh Epi-CFU medium three times weekly. 11. Visualize colony morphology by bright- or dark-field microscopy (Fig. 2G.1.2C-E). The fractionation strategy described in Basic Protocol 1 enriches for a population of cells containing both lineage-restricted airway and alveolar CFU and mixed-lineage CFU, which can be identified based on colony morphology (Fig. 2G.1.2D,E).

EXPANSION OF LUNG EPITHELIAL STEM/PROGENITOR CELLS IN CULTURE

BASIC PROTOCOL 3

In this protocol, EpiSPC are seeded in a stromal-free Matrigel culture supplemented with FGF-10 and HGF (Fig. 2G.1.3), in which they generate spherical cystic colonies that can be enzymatically dissociated and passaged weekly to maintain EpiSPC. This assay can also be used to assess the self-renewal capacity of putative EpiSPC. Somatic Stem Cells

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Figure 2G.1.3

Schematic representation of stromal-free culture setup for EpiSPC expansion.

NOTE: All procedures are performed in a sterile class II biological hazard flow hood or a laminar-flow hood. All solutions, reagents, media and equipment used to process and culture EpiSPC must be sterile, and proper aseptic technique should be used.

Materials Epi-CFU expansion medium (see recipe) Matrigel (standard concentration; BD Biosciences) Adult lung EpiSPC (CD45neg CD31neg EpCAMhi CD24low ) cells (Basic Protocol 1) Phosphate buffered saline (PBS; Sigma, cat. no. P-3813) Enzymatic digestion cocktail (see recipe) Refrigerated centrifuge Millicell-CM culture inserts (0.4-μm membrane, 12-mm diameter, hydrophilic PTFE, Millipore) 24-well flat-bottom tissue culture plates Triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ), humidified 15-ml conical tubes (e.g., BD Falcon) and 2-ml microcentrifuge tubes 21-G needles Set up expansion culture of lung EpiSPC 1. Following flow cytometric cell sorting, wash the sorted EpiSPC and MSC in 5 ml/tube Epi-CFU medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. 2. Resuspend individual cell pellets in Epi-CFU medium (1 to 2 ml per lung) and take an aliquot to determine cell concentration (UNIT 1C.3). Cell counts can be performed using an automated cell counter or a hemacytometer (UNIT 1C.3).

3. After determining cell concentrations, take an aliquot of Epi-SPC so that the final suspension contains the desired number of cells for culture (1000 cells per well). Allow for 100 μl per well. 4. Centrifuge EpiSPC cell suspension 5 min at 400 × g, 4◦ C, and aspirate medium. 5. Resuspend cell pellet in Matrigel diluted 1:1 with Epi-CFU expansion medium. Allow 100 μl per well. Ensure matrix always stays on ice otherwise it will solidify. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

6. Gently mix the Matrigel cell suspension. It is important to avoid creating bubbles. This can be achieved by holding the tube in the center of a Vortex mixer to create a swirling motion.

7. Place 12-mm Millicell-CM inserts in 24-well culture plates.

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8. Add 90 μl of Matrigel cell suspension atop of the filter membrane of a Millicell-CM insert and place plate in incubator (37◦ C) for 5 min to allow matrix to set. Be careful to avoid creating bubbles in the suspension.

9. Remove plate from incubator and add 400 μl of Epi-CFU expansion medium per well around the insert. 400 μl is just enough to allow the medium to touch the bottom of the insert, allowing diffusion of medium into the Matrigel without submerging the Matrigel culture.

10. Incubate cell cultures in a humidified 37◦ C triple–mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ) and change to fresh Epi-CFU expansion medium three times weekly.

Passage cells 11. After 1 week in culture, aspirate medium and wash cultures twice, each time with 1 ml sterile PBS. It is important to remove serum-supplemented medium because the serum will inhibit subsequent enzymatic digestion.

12. Harvest epithelial CFU by adding 1 ml of enzymatic digestion cocktail to the top of the insert and break up Matrigel by trituration. If clonal passaging is required, single colonies can be picked from the Matrigel and enzymatically digested rather than the bulk culture.

13. After Matrigel has been displaced from the insert, place enzymatic digestion cocktail (containing Matrigel and colonies) in a 2-ml microcentrifuge tube and incubate at 37◦ C for 30 min. 14. Using a 21-G needle, triturate the enzymatic digest to prepare a single-cell suspension. 15. Wash twice, each time with 5 ml Epi-CFU expansion medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. 16. Re-seed single-cell suspension by repeating steps 5 to 10. Cells should be passaged weekly and can be split into multiple cultures to accommodate the increase in cell concentration. The number of cells re-seeded depends on the progressive enrichment of colony-forming cells after sequential passage.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Enzymatic digestion cocktail PBS (tissue-culture grade; Invitrogen, cat. no. 14040) containing: 3 mg/ml collagenase (Type I) 3 mg/ml dispase Preheat at 37◦ C and use immediately Epi-CFU expansion medium Epi-CFU medium (see recipe) containing: 50 ng/ml recombinant FGF-10 (R&D Systems, cat. no. 345-FG) 30 ng/ml recombinant HGF (R&D Systems, cat. no. 2207-HG) Store up to 1 week at 4◦ C

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Epi-CFU medium α-MEM (Invitrogen, cat. no. 41061) containing: 10% (v/v) fetal bovine serum (heat inactivated) 1× penicillin/streptomycin (add from 100× stock) 1× insulin/transferrin/selenium (Invitrogen; add from 100× stock) 2 mM L-glutamine 0.0002% (w/v) heparin [1/1000 dilution of 0.2% heparin sodium salt (Invitrogen, cat. no. 07980) in PBS (Invitrogen, cat. no. 14040)] Store up to 4 weeks at 4◦ C Nycoprep 1.077 g/cm3 , 265 mOsm Combine the following: 300 ml Nycoprep Universal: (Nycodenz: 60% w/v solution), ready-made, sterile, endotoxin-tested, density = 1.310 g/cm3 ; 580 mOsm; 300 ml (Axis-Shield; http://www.axis-shield.com/)

300 ml sterile Tricine-NaOH (20 mM, pH = 7.2) 676.6 ml sterile 0.65% NaCl (w/v) Density = 1.077 gm/cm3 , Osmolarity = 265 mOsm, pH = 6.9 Store at room temperature and use before manufacturer’s expiration date COMMENTARY Background Information

Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

Identification and characterization of adult lung EpiSPC have been confounded by a lack of specific markers and functional assays for their prospective isolation, enumeration, and measurement of their proliferative and differentiative potential (Weiss et al., 2008; Chen et al., 2009; Bertoncello and McQualter, 2010). A number of studies have utilized flow cytometry for isolation of candidate lung stem/progenitor cell populations, including those based on the efflux of Hoechst 33342 which has proven to be selective for enriching Sca-1pos mesenchymal stromal cells (MSC; Reynolds et al., 2007; Summer et al., 2007). We have also demonstrated that sorting directly on the basis of Sca-1 expression enriches for MSC (McQualter et al., 2009). On the other hand, Kim et al. (2005) have reported a fractionation strategy in which sorting based on the co-expression of Sca-1 and CD34 resulted in the enrichment of a candidate CCSPpos Pro-SPCpos bronchioalveolar stem cell (BASC) cell subpopulation that retained epithelial character after serial passage in vitro. This protocol describes a strategy for isolating CD45neg CD31neg EpCAMhi CD24low lung EpiSPC, which are Sca-1low and clearly distinct from CD45neg CD31neg EpCAMneg Sca-1hi MSC or BASC (McQualter et al., 2010). The lack of concordance in the properties of cells isolated in these studies could

be explained by technical differences in tissue disaggregation (Raiser and Kim, 2009) and the limitations of in vitro assays used to assess proliferative and differentiative potential. The lack of knowledge of the intricate interactions between epithelial cells, mesenchymal cells, and the extracellular matrix has proven a significant obstacle in recapitulating the necessary conditions in vitro required for the development of assays for the identification of EpiSPC and the analysis of their organization and regulation. The culture system described in this unit utilizes a three-dimensional extracellular matrix (Matrigel), which allows the formation of a basement membrane for epithelial cell polarization and lumen formation, and enables the organotypic rearrangement of cells in culture recapitulating the physiological microenvironment of the lung. For that reason, this culture system can also be used to study epithelial-mesenchymal interactions that are important for lung regeneration and repair.

Critical Parameters To correct for spectral overlap between different fluorochromes during multicolor flow cytometric analysis and sorting, color compensation must be performed to correctly quantify the fluorescence intensity of each fluorochrome with which cells are labeled. When setting the level of compensation using cells from dissociated lung tissue, it is important to

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Figure 2G.1.4 Bivariate dot plot with back gating of EpiSPC (red) and MSC (blue) onto CD45neg CD31neg cells (gray) showing the potential overlap of these populations when plotted as CD45 + CD31 versus Sca-1.

take into account the autofluorescence of these cells. Alignment of autofluorescent cells with nonautofluorescent cells will result in overor under-compensation and inaccurate assignment of cell phenotype. Correct compensation assumes that the positive and negative populations have equal autofluorescence (Alexander et al., 2009; Alvarez et al., 2010). When applying the gating strategy used to sort cells by flow cytometry it is important to understand that very rarely are the boundaries between populations absolute. We have previously isolated lung MSC based on the CD45neg CD31neg Sca-1pos phenotype, which also comprised a minor population of epithelial CFU (McQualter, et al., 2009). However, subsequent introduction of EpCAM to the sort strategy demonstrated that all epithelial CFU could be removed from the MSC fraction by gating on the CD45neg CD31neg EpCAMneg Sca-1hi phenotype. Figure 2G.1.4 shows that when EpiSPC and MSC are back-gated onto our traditional plot of CD45+CD31 versus Sca1, the two subsets marginally overlap, which would explain why gates previously set for MSC using the CD45neg CD31neg Sca-1pos phenotype may also include a minor fraction of EpiSPC (McQualter et al., 2010). This protocol describes a method in which EpiSPC are cultured within Matrigel atop a Millicell-CM insert with medium supplied only at the basal surface to allow diffusion of nutrients into the Matrigel culture. It is important that the Matrigel layer not be submerged by medium, as this prevents colony formation. The Millicell-CM inserts chosen for this assay contain a special Biopore membrane (hydrophilic PTFE), which helps limit overgrowth of the stromal layer, and is transparent, allowing microscopic visualization.

The Epi-CFU described in these protocols have been grown under physiological low oxygen tension (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ), which has been shown to be optimal for growth of stem/progenitor cells at clonal density in vitro (Wion et al., 2009). However, Epi-CFU can be grown under standard oxygen tension (10% v/v CO2 in air), but the cloning efficiency may be lower.

Troubleshooting It is the authors’ experience that the differentiation state of MSC used in organotypic co-cultures is critical for supporting the growth of EpiSPC. It is important to use fresh CD45neg CD31neg EpCAMneg Sca-1hi MSC, as expansion of these cells in culture results in their differentiation and inhibits their ability to support growth of EpiSPC (manuscript in preparation).

Anticipated Results Using this protocol, the cell CD45neg CD31neg EpCAMhi CD24low fraction isolated comprises an enriched but heterogeneous population of lineagerestricted (airway or alveolar) epithelial progenitors and multipotent (multi-lineage) stem cells (∼2000 cells per lung), while the CD45neg CD31neg EpCAMneg Sca-1hi cell fraction represents a population of enriched MSC (∼100,000 cells per lung; McQualter et al., 2010). EpiSPC are cultured using two different techniques. Morphological characterization of colonies generated from EpiSPC grown in organotypic differentiation cultures demonstrates the generation of large lobular cystic colonies with a clearly defined lumen (airway-CFU), small dense saccular colonies

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(alveolar-CFU), and colonies of mixed phenotype with distinct budding (multipotentCFU, Fig. 2G.1.2). Under these conditions, MSC differentiate into lipid-filled fibroblasts and myofibroblasts. In stromal-free expansion cultures, EpiSPC generate smaller spherical colonies, which can be enzymatically passaged on a weekly basis.

Time Considerations Temporal analysis of colony formation in this organotypic assay system (Basic Protocol 2) results in the emergence of colonies after 5 days in culture, and their continued expansion and differentiation over a 2-week period. In stromal-free expansion cultures (Basic Protocol 3), optimal colony formation for serial propagation and re-seeding of CFU is achieved when colonies are harvested after 1 week. After this point, colonies begin differentiate and deteriorate, and the recloning efficiency of dissociated colonies is substantially reduced.

Literature Cited Alexander, C.M., Puchalski, J., Klos, K.S., Badders, N., Ailles, L., Kim, C.F., Dirks, P., and Smalley, M.J. 2009. Separating stem cells by flow cytometry: Reducing variability for solid tissues. Cell Stem Cell 5:579-583.

Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Kim, C.F., Jackson, E.L., Woolfenden, A.E., Lawrence, S., Babar, I., Vogel, S., Crowley, D., Bronson, R.T., and Jacks, T. 2005. Identification of bronchioalveolar stem cells in normal lung and lung cancer. Cell 121:823-835. McQualter, J.L., Brouard, N., Williams, B., Baird, B.N., Sims-Lucas, S., Yuen, K., Nilsson, S.K., Simmons, P.J., and Bertoncello, I. 2009. Endogenous fibroblastic progenitor cells in the adult mouse lung are highly enriched in the sca-1 positive cell fraction. Stem Cells 27:623633. McQualter, J.L., Yuen, K., Williams, B., and Bertoncello, I. 2010. Evidence of an epithelial stem/progenitor cell hierarchy in the adult mouse lung. Proc. Natl. Acad. Sci. U.S.A. 167:1414-1419. Raiser, D.M. and Kim, C.F. 2009. Commentary: Sca-1 and cells of the lung: A matter of different sorts. Stem Cells 27:606-611. Reynolds, S.D., Shen, H., Reynolds, P.R., Betsuyaku, T., Pilewski, J.M., Gambelli, F., Di Giuseppe, M., Ortiz, L.A., and Stripp, B.R. 2007. Molecular and functional properties of lung SP cells. Am. J. Physiol. Lung Cell Mol. Physiol. 292:L972-L983. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P, Orfao, A., Rabinovitch, P., and Watkins, S. 2010. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J.

Alvarez, D.F., Helm, K., Degregori, J., Roederer, M., and Majka, S. 2010. Publishing flow cytometry data. Am. J. Physiol. Lung Cell Mol. Physiol. 298:L127-L130.

Summer, R., Fitzsimmons, K., Dwyer, D., Murphy, J., and Fine, A. 2007. Isolation of an adult mouse lung mesenchymal progenitor cell population. Am. J. Respir. Cell Mol. Biol. 37:152159.

Bertoncello, I. and McQualter, J.L. 2010. Endogenous lung stem cells: What is their potential for use in regenerative medicine? Expert Rev. Respir. Med. 4:349-362.

Weiss, D.J., Kolls, J.K., Ortiz, L.A., PanoskaltsisMortari, A., and Prockop, D.J. 2008. Stem cells and cell therapies in lung biology and lung diseases. Proc. Am. Thorac. Soc. 5:637-667.

Chen, H., Matsumoto, K., and Stripp, B.R. 2009. Bronchiolar progenitor cells. Proc. Am. Thorac. Soc. 6:602-606.

Wion, D., Christen, T., Barbier, E.L., and Coles, J.A. 2009. PO(2) matters in stem cell culture. Cell Stem Cell 5:242-243.

Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

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Colon Cancer Stem Cells 1

UNIT 3.1 2, 3

Antonija Kreso and Catherine Adell O’Brien 1

Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada 2 Institute of Medical Sciences, University of Toronto, Toronto, Ontario, Canada 3 Department of Surgery, Division of General Surgery, University Health Network, Toronto, Ontario, Canada

ABSTRACT This unit describes protocols for working with colon cancer stem cells. To work with these cells one must start by generating single-cell suspensions from human colon cancer tissue. These cell suspensions are sorted using flow cytometry–assisted cell sorting to fractionate the cells into tumor-initiating and nontumor-initiating subsets. Once the cells have been fractionated, they must be functionally tested to determine tumor-forming capacity, the gold standard being the in vivo xenograft assay. Methods have also been developed to grow these cells in vitro in a sphere-forming assay. This unit will describe how to isolate and functionally test colon cancer stem cells, as well as provide advice on the potential challenges of the research. Curr. Protoc. Stem Cell Biol. 7:3.1.1-3.1.12. C 2008 by John Wiley & Sons, Inc.  Keywords: human colon cancer r cancer stem cells r in vivo xenograft assay r in vitro sphere assay

INTRODUCTION This unit describes protocols for working with colon cancer stem cells (CSC). The ability to successfully carry out this work is dependent on obtaining fresh colon cancer specimens at the time of surgical resection. Tissue fragments are processed to generate a single-cell suspension, which can then be fractionated utilizing flow cytometry to isolate subpopulations based on differential expression of cell surface markers, such as CD133 (O’Brien et al., 2007; Ricci-Vitiani et al., 2007; Todaro et al., 2007). Once these cell subsets have been fractionated, they can be tested for their tumor-forming capacity using the in vivo NOD/SCID xenograft assay. Utilizing this model it has been shown that tumor-initiating capacity exists solely within the CD133+ cell subset of colon cancer cells. The focus of this unit will be to describe the protocols for isolating, culturing (Basic Protocol 1), fractionating (Basic Protocol 2), and establishing a NOD/SCID xenograft model (Basic Protocol 3) to study colon CSC. The sphere-forming assay is also described (Basic Protocol 4). NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator. NOTE: All experiments using human tissue must be approved by the institutional committee on the ethical use of human subjects/material and tissue samples must be obtained with prior informed consent. Cancer Stem Cells Current Protocols in Stem Cell Biology 3.1.1-3.1.12 Published online November 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0301s7 C 2008 John Wiley & Sons, Inc. Copyright 

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NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals. BASIC PROTOCOL 1

GENERATING SINGLE-CELL SUSPENSIONS FROM HUMAN COLON CANCER TISSUE The first step to working with colon CSC involves generating a single-cell suspension from human colon cancer tissue. The percentage of necrotic cells in human tumors varies extensively and is dependent on multiple factors including: tumor characteristics, preoperative adjuvant chemotherapy or radiation therapy, and length of operative procedure. One factor that can help in obtaining the maximum number of viable cells is to ensure that specimens are received from the operating room expeditiously after removal from the patient. It has been our experience that each sample possesses a CSC fraction; however, the percent of this fraction can vary widely between tumors and this is true whether one is using CD133 or CD44 to isolate the CSCs. It is best when isolating the CSC fraction to start with at least 1 to 2 × 106 colon cancer cells (use a tumor fragment ∼1 × 0.5–cm in size to generate this many cells), because this will help ensure that there are enough cells in the CSC and non-CSC subsets to carry out the experiments.

Materials Colon tumor fragment Colon cancer stem cell medium (SCM; see recipe) Collagenase IV solution (200 U/ml SCM) Ammonium chloride: 0.8% (w/v) NH4 Cl in 0.1 mM EDTA Trypan blue 35-mm petri dishes Razor blade and forceps 5-ml disposable pipets 50-ml conical tube 45-μm cell filter Plunger from a 3- to 5-ml syringe Hemacytometer Additional reagents and equipment for counting cells using a hemacytometer and trypan blue (UNIT 1C.3) Isolate colon cancer cells 1. Place colon tumor fragment in 2 to 3 ml SCM in a 35-mm petri dish. 2. Using a razor blade and forceps, mince the tissue as much as possible. 3. Pipet tumor solution up and down 3 to 5 min with a 5-ml disposable pipet. Place the solution into a 50-ml conical tube. If fragments are too large to be drawn up into a 5-ml pipet, use a pipet with an opening large enough to draw up all the tumor fragments. Note that the highest cell numbers are typically obtained when tumors are minced to yield very small pieces.

4. Add the collagenase solution to the tumor cells. Incubate 30 to 60 min at 37◦ C. Pipet up and down a few times every 15 min. The final concentration should be 200 U of collagenase IV per milliliter of SCM.

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5. Pass the tumor solution through a 45-μm filter. Use a plunger from a 3- to 5-ml syringe and gently mash the tumor pieces to enable more tumor cells to pass through. Wash the filter with 4 to 5 ml of SCM.

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The tumors can be very different; some are soft whereas others are hard and fibrotic. The fibrotic tumors may not completely dissolve with collagenase. The fragments that remain on the filter can be resuspended in SCM with collagenase and incubated for 1 to 2 hr at 37o C. Then repeat step 5.

6. Centrifuge the tumor cell suspension 10 min at 450 × g, 4◦ C. 7. Resuspend the pellet in ∼5 ml of ammonium chloride (0.8% w/v NH4 Cl with 0.1 mM EDTA). Leave for 10 min at room temperature to lyse the red blood cells. After 10 min add an equal volume of SCM and centrifuge 10 min at 450 × g, 4◦ C. 8. Resuspend the pellet in 10 ml SCM. If the solution appears clumpy then pass it through another 45-μm filter. 9. Count an aliquot of the cells using a hemacytometer and trypan blue (UNIT 1C.3) to determine the percentage of dead cells. If there is a high percentage of necrotic cells, a Ficoll column can be used to remove the dead cells and debris (see Support Protocol).

USING A FICOLL COLUMN TO REMOVE DEAD CELLS The high percentage of necrotic cells and debris in some samples makes it exceedingly difficult to successfully carry out techniques such as flow cytometry–assisted cell sorting and transduction. In cases where samples have >30% dead cells this Ficoll protocol can allow for an enrichment of viable cells.

SUPPORT PROTOCOL

Materials Ficoll Colon cancer cell suspension (Basic Protocol 1) Colon cancer stem cell medium (SCM; see recipe) 15-ml conical tubes 5-ml pipet 1. Place 5 ml Ficoll into a 15-ml conical tube. 2. Resuspend the colon cancer cells in 5 ml of SCM. Layer this solution on top of the 5 ml of Ficoll. Divide the tumor cell suspension such that each 5 ml of medium contains no more than 4 to 5 × 106 tumor cells. If the cell number is >5 × 106 , divide the sample into the appropriate number of Ficoll-containing tubes.

3. Centrifuge 15 min at 1000 × g, 4◦ C. 4. Use a 5-ml pipet to remove 2 to 3 ml of medium off the top and then place the pipet at the interface (between the medium and Ficoll). Collect the interface, remainder of the medium, and a small amount of Ficoll. Resuspend the pellet in 5 ml SCM and save until the viable cell count is complete. Keep this solution until the cell number from the viable fraction has been counted. If the number of viable cells post-Ficoll differs significantly from the pre-Ficoll count, it is possible that some viable cells are in the pellet. In that case, repeat step 2 with the resuspended pellet.

5. Centrifuge 10 min at 450 × g, 4◦ C. Resuspend the pellet in a desired volume of SCM. Cancer Stem Cells

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BASIC PROTOCOL 2

FLOW CYTOMETRY–ASSISTED CELL SORTING Flow cytometric–assisted cell sorting is an essential aspect of CSC work. It is required to fractionate the CD133+ and CD133− cell subsets. It also allows the researcher to exclude all murine cells when sorting colon cancer xenografts, thereby avoiding any murine hematopoietic or endothelial cells contaminating the post-sort cell populations. To avoid contaminating cells in primary human colon cancer sorts one can positively select for epithelial specific antigen (ESA) expression and sort the following populations: ESA+ CD133+ and ESA+ CD133− . The initial selection on ESA+ cells allows one to exclude contaminating hematopoietic, endothelial, and stromal cells. Furthermore, flow cytometry also allows one to study other markers of interest in combination with CD133. If flow cytometry–assisted cell sorting is not available, another option is to carry out magnetic bead cell sorting as per the Miltenyi-Biotec protocol. It has been our experience that this method can be used successfully provided the starting sample has ≤30% dead cells. If the sample has >30% dead cells, it can be difficult to obtain the necessary purity (≥90% to 95%) using the MACS bead separation. To successfully carry out MACS bead enrichment the sample should be passed through at least three Miltenyi-Biotec columns in order to best enrich the sample. One of the other main disadvantages of using the MACS beads is the inability to select on multiple cell surface markers.

Materials Colon cancer cell suspension with <30% dead cells Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) with 0.1% bovine serum albumin (BSA; CMF-PBS/0.1% BSA) ESA antibody conjugated to a fluorophore Anti–mouse antibody conjugated to a fluorophore CD133 APC or PE (Miltenyi Biotec) Propidium iodide (final concentration: 1 μg/ml of PBS with 0.1% BSA) 5-ml polystyrene tubes Additional reagents and equipment for flow cytometry–assisted cell sorting (Robinson et al., 2008) and counting cells using a hemacytometer (UNIT 1C.3) 1. Resuspend colon cancer cells in PBS/0.1% BSA to a concentration of 0.5 to 1 × 106 cells/100 μl. During antibody staining, if you have ≤5 × 106 cells, you can stain all cells in the 100 μl per 1 × 106 cells. However, if you have >5 × 106 , divide the cells into separate polystyrene tubes for staining. This is done because it is best to stain cells in a smaller volume.

2. Add fluorophore-conjugated anti-CD133 and anti-mouse or anti-ESA antibodies to the sort sample. When sorting human samples, an antibody against ESA should be used. When sorting xenografts, use an anti-murine IgG antibody to exclude murine cells. Check with the FACS facility to determine whether isotypes and individual antibody control stains are required. All antibodies should be titrated to determine the required amount. Ask the FACS facility whether they prefer that the sort sample be placed into a polypropylene or polystyrene tube for sorting.

3. Incubate cells with antibodies 30 to 45 min at 4◦ C, protected from light. 4. Add 4.5 ml CMF-PBS/0.1% BSA to each tube and centrifuge 5 min at 450 × g, 4◦ C. Discard the supernatant. Repeat three times to wash the cells. Colon Cancer Stem Cells

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5. Following the last wash, resuspend the cells in 1 to 2 ml CMF-PBS/0.1% BSA that contains 1 μg/ml propidium iodide. At this point the cells are ready to be sorted. Propidium iodide stains dead cells thereby allowing them to be excluded.

6. Sort the ESA+ CD133+ cells (Robinson et al., 2008) into 2 ml SCM. 7. Count an aliquot of the post-sort cells with a hemacytometer (UNIT 1C.3) to confirm the cell yield. The flow cytometry facility will provide a post-sort cell count for each population; however, it is important that the researcher confirm the cell count.

IN VIVO XENOGRAFT ASSAY The utilization of animal models is crucial in CSC work. The ideal choice is always an orthotopic model in which cancer cells are injected into the same tissue from which they are derived. However, when carrying out CSC research the most important factor is to identify an animal model that has the greatest reliability for xenograft formation. This is essential because if the tumor take rate for the model you choose is only 50% it becomes impossible to determine whether the absence of a xenograft is due to the lack of a CSC or simply the limitation of the animal model. Published colon CSC work to date have used two models: subcutaneous (Ricci-Vitiani et al., 2007; Todaro et al., 2007) and subrenal capsule (O’Brien et al., 2007) injections. It has been our experience that only ∼30% of colon cancer cell suspensions have a reliable take rate in the subcutaneous site. In our hands, injection of colon cancer cell suspensions under the renal capsule provided the most reliable results with almost all tumors tested giving rise to xenograft formation (overall take rate of ∼90%).

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Aside from the choice of injection site a decision must also be made about the type of immunocompromised mouse to be used. Published studies in colon CSC research have used either NOD/SCID or SCID mice (Dalerba et al., 2007; O’Brien et al., 2007; RicciVitiani et al., 2007). It is difficult to make a direct comparison between the efficiency of the NOD/SCID versus SCID mice for use in CSC work because no head-to-head comparison has been carried out between the two strains using the same marker set and injection site. It is also important to acknowledge that an increasing number of immunocompromised mouse strains are becoming available and may represent new options for carrying out this work. Irradiating the mice prior to the procedure can also improve the xenograft take rate; however, the marginal improvement in xenograft formation must be weighed against the radiation sensitivity of NOD/SCID mice. Irradiation should be carried out the day before or the day of the procedure and the dose should be 300 rad. It is best to carry out a trial of irradiation to determine both the potential benefit to xenograft take rate and the radiation sensitivity of the mice in the colony. Death related to radiation sensitivity usually occurs 6 to 7 weeks post-irradiation.

Materials Matrigel ∼10 μl sorted cell suspension in SCM (Basic Protocol 2) Stem cell medium (SCM; see recipe) NOD/SCID or SCID mice (8- to 10-week-old) Iodine-based solution (e.g., Betadine) 70% ethanol

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Normal saline, sterile Pain medication (e.g., buprenorphine) 1-ml syringe without needle 1-ml insulin syringes with a 29-G needle, 1/2 -in. long Heating pad Clippers Sterile gauze Scissors Forceps Sutures or surgical clips (Roboz) Additional reagents and equipment for rodent anesthesia using isoflurane (UNIT 1B.4) 1. Thaw an aliquot of Matrigel and draw it up into a 1-ml syringe without needle. Then remove the plunger from a 1-ml insulin syringe with a 29-G needle, 1/2 -in. long and insert a small amount of Matrigel into the back of the syringe (Fig. 3.1.1A). It is difficult to be exact on the amount of Matrigel—one should aim for it to be 25 to 50 μl (closer to 25 μl is better). The best way to estimate the amount of Matrigel is to look at the markings on the side of the syringe. Keep undiluted Matrigel aliquoted and frozen at −20o C, ∼100 to 200 μl per microcentrifuge tube.

2. Use a pipet to inject the cell suspension (aim to resuspend the cells in 10 μl of SCM) into the middle of the Matrigel (Fig. 3.1.1B). Then reinsert the plunger into the syringe and push the mixture to the top of the syringe (Fig. 3.1.1C,D). Once the syringes are prepared they should be injected in a timely fashion. It is best to keep the syringes on ice, because at room temperature Matrigel will set.

Figure 3.1.1 Preparing syringes for injection. (A) Inserting the Matrigel into the back of the insulin syringe. (B) Using a pipet place the 10 μl of cell solution into the middle of the Matrigel. (C) Reinsert the insulin syringe plunger. (D) Using the plunger push the Matrigel and cell solution to the top of the syringe, it is now ready for injection. Colon Cancer Stem Cells

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Figure 3.1.2 Subrenal capsule injection. (A) The mouse is positioned with its left side up. A left flank incision is made just under the costal margin. (B) The kidney is gently pulled out of the abdominal cavity, being careful not to disturb the blood supply. (C) The needle is positioned just under the renal capsule. (D) Following the injection of Matrigel and cell suspension there is a small bleb on the surface of the kidney.

3. Handle mice using sterile technique and anesthetize using inhalational anesthesia (UNIT 1B.4). Place the mice on a heating pad during the procedure. 4. Position the mouse left side up. Use clippers to shave the area ∼0.5-cm below the costal margin on the left side. Wash the clipped area sequentially with iodine-based solution and 70% ethanol solution and then dab dry with sterile gauze. 5. Using scissors make an ∼0.5-cm long incision on the flank, just below the costal margin on the left side (Fig. 3.1.2A). Only inject into the left kidney. Due to the anatomy it is very difficult to inject into the right kidney. Cancer Stem Cells

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6. Gently pull the kidney from the abdominal cavity using two pairs of forceps. Then take the syringe containing the cells to be injected and place the needle just under the kidney capsule and push the needle until just before it comes out the opposite pole. At this point, inject the cells as you slowly pull back on the needle. The cell solution should be completely injected before the needle exits the kidney (Fig. 3.1.2B,C,D).

7. Deliver the kidney back into the abdomen and close the abdominal wall. There is the choice to close using sutures or surgical clips (Roboz) both of which provide equivalent results. The choice may depend on the requirements of the animal research facility at your particular institution.

8. Prior to awakening the mouse from anesthesia administer a 1 ml subcutaneous bolus of sterile normal saline and a dose of pain medication (e.g., 0.01 to 0.05 mg/kg buprenorphine). At the commencement of this work an antibiotic (Baytril) was added to the drinking water of all mice post-procedure for 2 weeks. The mice became dehydrated and as a result weak; therefore, the practice was stopped. There were no deleterious effects from discontinuing Baytril.

9. Assess the mice for tumor development every week starting 2 weeks post procedure. This can be done by holding the mouse and gently palpating in the area of the kidney. Over time, you will start to appreciate a fullness (the tumor will feel like a firm nodule) in this area. The rate of tumor development will differ depending on the tumor. Some xenografts will appear in 6 to 8 weeks; however, others can take up to 30 weeks to develop. BASIC PROTOCOL 4

CULTURING COLON CANCER CELLS AS SPHERES The ability to culture colon CSC requires the utilization of a serum-free stem cell medium. Using this protocol colon CSC will grow as spheres (Fig. 3.1.3A,B) in a non-adherent manner. The addition of serum to this medium results in the differentiation of the colon cancer cells and their growth as an adherent layer. Although much of the current work in the cancer stem cell field has been carried out using in vivo models, the ability to culture the cells as spheres can be used to complement the in vivo work. It is important to keep a record of the passage number for each tumor in vitro.

Materials Colon cancer cell suspension, sorted (Basic Protocol 2) Stem cell medium (SCM; see recipe) Trypsin/EDTA Ultra-low attachment surface dishes (Corning) 5-ml disposable pipet 45-μm filter Additional reagents and equipment for counting cells using trypan blue (UNIT 1C.3) 1. Plate sorted colon cancer cells at a density of 30,000 to 50,000 cells/ml of SCM. For best results the spheres should be grown in ultra-low attachment surface dishes (Corning). Any size dish can be used depending on the cell number; the most important point is to plate at a density of 30,000 to 50,000 cells/ml of SCM.

2. Passage the cells approximately every 4 days. Colon Cancer Stem Cells

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There is some variability between tumors and, therefore, each tumor must be evaluated daily to follow sphere formation. Passaging requires the disruption of the spheres to generate a single-cell suspension. There are two possible approaches to sphere disruption: mechanical or enzymatic. Current Protocols in Stem Cell Biology

Figure 3.1.3 Photographs of colon cancer sphere cultures. (A) Three colon cancer spheres (magnification 4×). (B) One colon cancer sphere (magnification 20×).

Mechanical sphere disruption 3a. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 4a. Resuspend the pellet in 3 ml of SCM and pipet up and down for 10 min with a 5-ml disposable pipet. After pipetting for 10 min look at the solution. There should no longer be any visible spheres. If visible spheres remain, continue to pipet for another 5 to 10 min.

5a. Pass solution through a 45-μm filter, count an aliquot of the cells (UNIT 1C.3), and resuspend in the desired volume of SCM for replating. Best results are obtained when cells are plated at a density of 30,000 to 50,000 cells/ml of SCM.

Enzymatic sphere disruption 3b. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 4b. Resuspend the pellet in 3 to 5 ml of 1× trypsin/EDTA and pipet up and down for 3 min with a 5-ml disposable pipet. At this point, place the tube in the incubator 10 min at 37◦ C. 5b. After 10 min remove the cells from the incubator and add an equal volume of SCM. Pass the cells through a 45-μm filter. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 6b. Resuspend the cell solution and count the number of viable cells using trypan blue (UNIT 1C.3). The criticism associated with the use of enzymatic digestion is that it may interfere with the expression of cell surface markers. Enzymatic digestion can be used; however, it should be tested initially against mechanically digested cells from the same tumor. This testing should be carried out for each cell surface marker to determine whether enzymatic digestion has any deleterious effect on the expression of the cell surface markers being studied.

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Growth factor mix, 10× For 200 ml of growth factor mix: 100 ml DMEM/F12 4 ml 30% (w/v) glucose 200 mg transferrin 50 mg insulin in 20 ml of water (add 2 ml of 0.1 N HCl to dissolve, then add 18 ml of water) 19.33 mg putrescine in 20 ml water 200 μl 0.3 mM sodium selenite 20 μl 2 mM progesterone H2 O to 200 ml Divide the growth factor mix into 2-ml aliquots and store indefinitely at −20◦ C Stem cell medium (SCM) 500 ml of a 1:1 ratio of DMEM/F12 (Invitrogen) 1% penicillin-streptomycin (1× from a 100× stock purchased from Invitrogen) 2 ml 50× B27 supplement (Invitrogen) 4 μg/ml heparin 1% (w/v) of non-essential amino acids 1% (w/v) of sodium pyruvate 1% (w/v) of L-glutamine Store up to 1 week at 4◦ C Just before use, add 5 ml growth factor mix (see recipe) 10 ng/ml fibroblast growth factor 20 ng/ml epidermal growth factor COMMENTARY Background Information

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The existence of a CSC fraction was first studied in the context of human leukemia. Lapidot et al. demonstrated that acute myelogenous leukemias possess a CSC subset capable of recapitulating the disease in a SCID mouse model, whereas the non-CSC cells were incapable of generating the disease (Lapidot et al., 1994). More recently, it has been shown that a wide variety of solid tumors also possess a CSC subset including breast (Al-Hajj et al., 2003), brain (Singh et al., 2004), and colon cancers (O’Brien et al., 2007; Ricci-Vitiani et al., 2007). The identification of a CSC population in human colon cancer was first published in 2007, when two groups established that fractionation of colon cancer cells based on CD133 expression identified a subset of CD133+ cancer cells that was capable of initiating tumor growth in murine xenograft models. In con-

trast, the CD133− cancer cells were unable to initiate tumor growth. The limiting dilution analyses in one of the studies demonstrated that ∼1 in 262 CD133+ colon cancer cells represented a CSC, for the ten tumors tested in the series, thereby demonstrating that CD133 expression enriches for tumor initiating capacity but does not identify a pure CSC population (O’Brien et al., 2007). More recently, another publication demonstrated that CD44 and CD166 (ALCAM) expression could also be utilized to enrich for a CSC subset in colon cancers (Dalerba et al., 2007). It is important to appreciate that the markers identified to date enrich for CSC; however, they do not identify a pure population. The field remains at a very nascent stage, and advancement will depend in large part on the identification of new CSC markers that can be used in conjunction with markers, such as CD133 and CD44, to provide further enrichment of the CSC fraction.

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Limiting dilution analysis (LDA) represents an essential tool in carrying out CSC work because it provides the ability to calculate the frequency of CSC within a population of cancer cells. LDAs require the injection of a range of doses with multiple mice being injected per dose (Porter and Berry, 1964). The gold standard is to carry out an LDA of both bulk and fractionated cancer cells for each individual tumor. This allows for the calculation of a CSC frequency in both the bulk tumor cell population and the fractionated subpopulations (e.g., CD133+ versus CD133− ). The limiting factor in these experiments is often the paucity of cell number. One method to circumvent issues of cell number is to initially inject unsorted cells into four to five mice to expand the cell number and then to use these xenografts to carry out the bulk and fractionated LDAs in mice. It has been our experience and the experience of others that the cell surface phenotype is maintained following passage in mice (Dalerba et al., 2007; O’Brien et al., 2007; Ricci-Vitiani et al., 2007). However, it is very important to determine the cell surface phenotype for each tumor prior to passage in mice and then after each passage. This will allow you to confirm for each tumor that the subpopulations are remaining stable following passage in vivo. It has been our experience that at high levels of in vivo passage (7 and above) we do start to see some tumors that change their cell surface phenotype; however, the changes are not predictable. It is for these reasons that it is crucial to check the tumor cell surface phenotype with each passage. There is very limited data on the propagation of colon CSC as sphere-forming units. One recent publication identified that in a series of colon cancers only approximately half could be propagated in vitro as spheres (Todaro et al., 2007). There is also the suggestion that CSC marker expression may change with in vitro propagation (A. Kreso and C.A. O’Brien, unpub. observ.). Therefore, the sphere-forming assay represents a surrogate; however, it does not eliminate the need to carry out the gold standard, functional in vivo assays. The sphere-forming assay requires further study to clearly establish its role within CSC work and to determine how closely it recapitulates in vivo models. Furthermore, caution must be exercised when using cancer cells following serial passages in vitro because to date it has not been clearly established whether the cells maintain the same functional phenotypes. Until these questions have been answered, the best approach when

using a sphere-forming assay is to functionally test the CSC and non-CSC fractions with in vivo assays at a minimum of every other in vitro passage.

Critical Parameters and Troubleshooting The ability to successfully sort the cells using flow cytometry will depend in large part on the flow facility. It is important when starting this work to determine the level of expertise at your flow facility for sorting solid tumor cells. If the facility does not have expertise in this area, it is important to contact a flow facility that regularly sorts solid tumor cells to establish the instrument settings that result in the highest yield, both with respect to purity and viability of the cells post-sorting. It is also essential to count cells post-sorting to confirm the cell yield. There can be a discrepancy between the stated and actual cell yields; having an accurate cell count is essential when carrying out LDA experiments.

Anticipated Results The in vivo protocol will result in the generation of xenografts, which recapitulate the phenotype of the original tumors. At the time of sacrifice a fragment of each xenograft should be saved for histological assessment. This will allow the researcher to confirm that the xenograft recapitulates the original tumor with respect to differentiation status and tumor subtype. The in vitro protocol will result in expansion of the colon cancer cells; however, as previously stated there is a proportion of the cancers that cannot be successfully passaged in vitro. At this time we are unable to predict which tumors will grow well in vitro. As more work is carried out in the field, it will hopefully lead to a better understanding of the in vitro requirements to maintain colon cancer cells in serum-free culture conditions.

Time Considerations The initial culturing of colon CSC from an unpassaged primary human specimen may take >4 to 5 days. If this is the case, one should leave the cells in the incubator for 7 to 14 days, adding EGF and FGF to the medium every 4 days. If the tumor has not formed spheres by 14 days it is unlikely to do so. The time for xenograft formation varies between tumors, ranging from 6 to 24 weeks. To do the in vivo experiments it is important to use NOD/SCID mice between the ages of 8 and 10 weeks. Mice utilized before 8 weeks

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have an increased chance of succumbing to the stress of the procedure. Using older mice (>10 weeks) can be difficult because as NOD/SCID mice age there is a natural attrition rate; therefore in the case of tumors that take 24 weeks to appear it is possible that the mice of interest will die before the appearance of xenografts. The primary tissue is often received from the operating room at the end of the workday and it is important to note that once the cell suspension is generated these cells can be safely left at 4◦ C overnight in SCM and injected or sorted the next day.

Acknowledgements We thank Sean McDermott for his contributions to the Ficoll protocol and other members of the Dick laboratory for helpful discussions. We also acknowledge members of Peter Dirks’ laboratory, especially Ian Clarke, for technical suggestions. We thank John E. Dick for his invaluable help and guidance.

Literature Cited Al-Hajj, M., Wicha, M.S., Benito-Hernandez, A., Morrison, S.J., and Clarke, M.F. 2003. Prospective identification of tumorigenic breast cancer cells. Proc. Natl. Acad. Sci. U.S.A. 100:39833988. Dalerba, P., Dylla, S.J., Park, I.K., Liu, R., Wang, X., Cho, R.W., Hoey, T., Gurney, A., Huang,

E.H., Simeone, D.M., Shelton, A.A., Parmiani, G., Castelli, C., and Clark, M.F. 2007. Phenotypic characterization of human colorectal cancer stem cells. Proc. Natl. Acad. Sci. U.S.A. 104:10158-10163. O’Brien, C.A., Pollett, A., Gallinger, S., and Dick, J.E. 2007. A human colon cancer cell capable of initiating growth in immunodeficient mice. Nature 445:106-110. Porter, E.H. and Berry, R.J. 1964. The efficient design of transplantable tumour assays. Br. J. Cancer 17:583-595. Ricci-Vitiani, L., Lombardi, D.G., Pilozzi, E., Biffoni, M., Todaro, M., Peschle, C., and DeMaria, R. 2007. Identification and expression of human colon-cancer-initiating cells. Nature 445:111-115. Robinson, J.P., Darzynkiewicz, Z., Dean, P.N., Dressler, L.G., Rabinovitch, P.S., Stewart, C.C., Tanke, H.J., and Wheeless, L.L. (eds.) 2008. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Singh, S.K., Hawkins, C., Clarke, I.D., Squire, J.A., Bayani, J., Hide, T., Henkelman, R.M., Cusimano, M.D., and Dirks, P.B. 2004. Identification of human brain tumor initiating cells. Nature 432:396-401. Todaro, M., Alea, M.P., Di Stefano, A.B., Cammareri, P., Vermeulen, L., Iovino, F., Tripodo, C., Russo, A., Gulotta, G., Medema, J.P., and Stassi, G. 2007. Colon cancer stem cells dictate tumor growth and resist cell death by production of interleukin-4. Cell Stem Cell 1:389-402.

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In Vivo Evaluation of Leukemic Stem Cells through the Xenotransplantation Model

UNIT 3.2

Dominique Bonnet1 1

Cancer Research UK, London Research Institute, Haematopoietic Stem Cell Laboratory, London, United Kingdom

ABSTRACT The xenotransplantation model has been instrumental for the identification and characterization of human leukemic stem cells. This unit describes our current method for the engraftment of human leukemic patients’ samples in the xenotransplanted mouse model. We concentrate uniquely on the model of acute myeloid leukemia, as it was the first type of leukemia for which the xenotransplantation model was developed. Nevertheless, the Basic Protocol could be applied to other sorts of blood disorders. Curr. Protoc. Stem Cell C 2008 by John Wiley & Sons, Inc. Biol. 7:3.2.1-3.2.11.  Keywords: hematopoietic stem cell (HSC) r xenotransplantation r immunodeficient mice r leukemic stem cell (LSC)

INTRODUCTION The adaptation of xenotransplantation assays to examine the propagation of acute myeloid leukemia (AML) in vivo has been fundamental in the identification and characterization of leukemia-initiating cells (Lapidot et al., 1994; Bonnet and Dick, 1997). Transplantation of primary AML cells into NOD/SCID mice led to the finding that only rare cells, termed AML-initiating cells (AML-IC), also known as leukemic stem cells (LSC), are capable of initiating and sustaining growth of the leukemic clone in vivo, and serial transplantation experiments showed that AML-IC possess high self-renewal capacity, and thus can be considered to be the leukemic stem cells. The Basic Protocol below describes the most common and simple method to test for the presence of leukemia-initiating cells; this method can also be used to characterize blood disorders. Support protocols describe methods for further purification of leukemic stem cells and the intra-bone injection procedure. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. NOTE: All of the following procedures using human samples should be performed in a level 2 safety tissue culture unit using sterile and proper aseptic techniques.

IDENTIFICATION OF LEUKEMIA STEM CELLS THROUGH XENOTRANSPLANTATION

BASIC PROTOCOL

Xenotransplantation of human cells obtained from AML patients allows identification and characterization of leukemic stem cells.

Materials Immunodeficient mice: NOD/SCID, NOD/SCID-β2 microglobulin null (β2m−/− ), or NOD/SCID IL2R gammanull (Jackson Laboratory) Acidified water: a solution of HCl at a final pH 2.8 to 3.2 Current Protocols in Stem Cell Biology 3.2.1-3.2.11 Published online December 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0302s7 C 2008 John Wiley & Sons, Inc. Copyright 

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AML sample: peripheral blood or bone marrow Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) Ammonium chloride solution (Stem Cell Technologies) Fetal bovine serum (FBS; Stem Cell Technologies, cat. no. 06471) Antibodies against human CD45, CD34, CD38, CD33, and CD19 (BD Biosciences Pharmingen) 100 ng/ml 4 ,6-diamidino-2-phenylindole (DAPI; UV excited, Sigma-Aldrich) or TOPRO-3 [HeNe (633-nm) excitable, Molecular Probes] Irradiator: Cesium source is recommended, but an X-ray system or Cobalt source can also be used 29-G, 1/2 -in. needle and insulin syringe (Tyco Healthcare) Dissection tools: scissors and forceps 5-ml snap-top polystyrene tubes Benchtop centrifuge equipped with swing-out bucket rotor for 15- and 50-ml conical tubes Hemacytometer Fluorescent-activated cell sorter, e.g., FACSAria (BD Biosciences) and/or a Moflow (Dako) equipped with 488-nm, 633-nm, and 404-nm lasers 440/40 bandpass (bp) filter for analysis of DAPI, a 530/30 bp filter for FITC, a 575/26 bp for PE, a 695/40 bp for PerCP, and a 660/20 bp for TOPRO-3 Mouse depletion kit (e.g., StemCell Technologies, cat. no. 13066) Additional reagents and equipment for assessing for AML engraftment (Support Protocol 3), parenteral injections (Donovan and Brown, 2006a), euthanasia of mice (Donovan and Brown, 2006b), and performing a cell count using a hemacytometer (UNIT 1C.3) Prepare the immunodeficient mice 1. Keep the animals in a pathogen-free environment. All the NOD/SCID animals used have some impairment of their immune system and may succumb to infections not affecting normal mice. They thus should be kept in pathogen-free status within barrier systems to protect them from current infections.

2. Treat the mice for at least 8 days with acidified water before irradiation. 3. Set the sublethal irradiation dose (between 300 and 375 cGys) on the irradiator used. Success indeed depends on the dose rate/minute of irradiation. If the dose rate/minute is initially too high, try to reduce it by using appropriate shielding. This will reduce the damage to internal organs.

4. Sublethally irradiate the mice before the adoptive transfer of cells. For best results, perform the irradiation 24 hr before the adoptive transfer of cells. The dose of irradiation depends on the mouse strains used and also on the source and irradiator used. It usually varies from 300 to 375 cG (see notes above).

5. Maintain any mouse receiving irradiation on acidified water for 2 weeks following the irradiation dose to prevent diarrhea or weight loss possibly arising due to epithelial damage of the intestines. Any animal showing persistent weight loss >20% of body weight and/or other signs of illness (e.g., rough fur, loss of appetite, inability to groom, and immobility) should be sacrificed. Experience shows that with these measures, the above side effects rarely arise.

Xenotransplantation Model of Leukemia

Prepare AML cells 6. Isolate mononuclear cells (for an example procedure, see Bonnet and Dick, 1997) from fresh peripheral blood or bone marrow samples. Frozen samples can also be used (see Support Protocol).

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7. For prescreening, inject 5 to 10 × 106 mononuclear cells per mouse (4 to 5 mice are tested). Not all AML samples at diagnosis engraft. Thus, for all new samples that arrive in the laboratory, we test the capacity to engraft first by injecting 5 to 10 × 106 viable cells/mouse using intravenous (tail vein) injections. Injection of the cells is done using under 100 to 200 μl/mouse. Usually, cells are resuspended in PBS/2% FBS.

8. After 8 to 12 weeks, sacrifice the mice by cervical dislocation (Donovan and Brown, 2006b) or terminal anesthesia and assess for AML engraftment (see Support Protocol 3). If the AML sample engrafts, purification of the AML-initiating cells (AML-IC) can be performed using Lin, CD34, and CD38 expression and cell sorting (see Support Protocol 3).

Transfer the cells adoptively 9. Subject mice to sublethal irradiation prior to injection of cell preparations (unpurified or purified cell fraction, genetically modified or not). This may be performed on the same day as, or up to 3 days after irradiation.

10. Inject between 106 and 107 cells intravenously via the tail vein (Donovan and Brown, 2006a; maximum volume 1% body weight) using a syringe with a 29-G, 1/2 -in. needle. In some cases, an intra-bone marrow injection (see Support Protocol) might be preferred, especially if a decrease in homing efficiency of the cell transferred is suspected.

Analyze the engraftment 11. Sacrifice mice between 4 to 14 weeks after transplantation using either cervical dislocation (Donovan and Brown, 2006b) or terminal anesthesia. When work under terminal anesthesia is involved, the level of anesthesia should be maintained at sufficient depth for the animal to feel no pain. Blood sampling is not informative as blood samples do not usually match the level of engraftment present in the bone marrow. Indeed, usually few AML cells circulate in the periphery, except in some AML samples (usually samples from patients with poor prognosis) where the AML infiltrates solid organs like spleen and liver. In these cases, the animals become sick and will need to be sacrificed potentially before 10 to 12 weeks.

12. Dissect the femurs, tibias, and iliac crests from the mice and store at room temperature in CMF-PBS before flushing. Remove all connective tissue around the bone.

Prepare bone marrow 13. Place 1 ml of room temperature CMF-PBS in a 5-ml snap-top polystyrene tube. This will be used to flush the bone marrow (see step 15, below).

14. Cut both ends of each bone to provide openings. 15. To flush, insert the 1-ml CMF-PBS-containing insulin syringe into one end of each bone and wash the lumen of the bone with medium pressure. Repeat twice for both ends of the bone or until the bone appears white.

Prepare the cells for FACS analysis 16. To lyse red blood cells, first cool the cell suspension 5 min on ice. Following the cooling of the suspension, add 3 ml of cold ammonium chloride solution to the 1-ml CMF-PBS/cell suspension, mix, and leave for 5 min at 4◦ C. 17. Add 0.5 ml FBS and centrifuge cells 5 min at 380 × g, 4◦ C. 18. Remove the supernatant and resuspend the cells in 1 ml of cold CMF-PBS containing 2% (v/v) FBS.

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Figure 3.2.1

(legend at right)

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19. Count the cells using a hemacytometer (UNIT 1C.3). Store on ice until ready for antibody labeling.

Stain the cells 20. Prepare a mix of human-specific FITC-conjugated anti-CD19, PE-conjugated antiCD33, and PerCP-conjugated anti-CD45 (5 μl/sample/antibody for all stains and compensation/isotype controls). Also, prepare FITC, PE, and PerCP single-color compensation control tubes (5-ml snap-top polystyrene tubes as before) and a combined FITC/PE/PerCP matched isotype control tube. 21. Distribute 15 μl of the antibody mix into each tube of a fresh set of 5-ml tubes for antibody labeling. 22. Dispense 40 μl of each cell suspension into the appropriate antibody labeling tube and leave to label for 30 min at 4◦ C. 23. Wash cells in 2 ml PBS/2% FBS and resuspend in 500 μl of PBS/2% FBS supplemented with a cell impermeant DNA dye for live/dead discrimination, either 100 ng/ml 4 ,6-diamidino-2-phenylindole or TOPRO-3.

Perform FACS analysis 24. To analyze this combination of fluorochromes set up the FACS device with a 488nm excitation source and either a UV or HeNe (633-nm) source depending on your choice of live/dead discriminator. 25. For emission collection ensure you have a 440/40 bandpass (bp) filter for analysis of DAPI, a 530/30 bp filter for FITC, a 575/26 bp for PE, a 695/40 bp for PerCP and a 660/20 bp for TOPRO-3 in place. 26. During FACS analysis, set the photomultiplier gains so that the background signal from the combined isotype control gives 1% to 5% positive cells in each collection channel. 27. Set the compensation amount according the detected spectral overlap. 28. To analyze the engraftment, draw four dotplots as in panels A, B, C, and D in Figure 3.2.1 [440/40 nm versus side-scatter (SSC), forward scatter (FSC) versus SSC, 695/40 nm versus SSC, and 530/30 nm versus 575/26 nm]. 29. First, exclude dead cells from the analysis via a region (R1) around the live, unstained cells as in Figure 3.2.1A. 30. Next, display these cells on a FSC versus SSC plot and select the lymphoid and myeloid cells for further analysis but exclude debris via a region (R2) as in Figure 3.2.1B. 31. Display cells that fall into the first two regions on a 695/40-nm versus SSC plot and draw a generous region around the CD45-PerCP positive cells as in Figure 3.2.1C (R3). Figure 3.2.1 (at left) For analysis of the engraftment by FACS, first the dead cells are excluded using DAPI staining and a live cells region (R1) is drawn (A). Next, these cells are displayed on an FSC versus SSC plot and lymphoid and myeloid cells are selected for further analysis, but debris is excluded via region R2 (B). A generous region is drawn (R3) around the CD45+ cells (C). These CD45+ cells are further analyzed for the expression of CD33 and CD19 (D). The number of events that fall within these regions may be used to calculate the percentage of live, debris-free cells (R2) that are human cells. In addition, the scatter characteristics of cells may be confirmed as consistent with myeloid [high FSC and SSC; (E)] and lymphoid [low FSC and SSC; (F)]. In panel F, there are no CD19+ cells. Engraftment is classed as myeloid leukemia if a population of CD45+ /CD33+ cells is present without an accompanying CD45+ CD19+ /CD33− cell population.

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32. Display these CD45+ cells on a CD19-FITC versus CD33-PE (530/30-nm versus 575/26-nm) dotplot and draw a quadrant to define FITC+ /PE− cells and FITC− /PE+ cell subsets as in Figure 3.2.1D. 33. Use the number of events that fall within these regions to calculate the percentage of live, debris-free cells (R2) that are human cells. In addition, evaluate the scatter characteristics of cells to confirm them as consistent with myeloid (high FSC and SSC, example in Fig. 3.2.1E) and lymphoid (low FSC and SSC, example in Fig. 3.2.1F). In this example, there are no CD19+ cells. Engraftment is classed as myeloid leukemia if a population of CD45+ /CD33+ cells is present without an accompanying CD45+ CD19+ /CD33− cell population. To confirm the leukemic origin of the myeloid cells present in the bone marrow of engrafted mice and if the original AML sample has a known translocation, it is possible to sort human CD45+ cells and perform fluorescent in situ hybridization analysis of the cells or any PCR analysis in search of a fusion product or a mutated gene (NPM, Flt3L, c-kit, WT1, CEBPα). The description of the procedures for FISH or PCR analysis is beyond the scope of this unit.

Perform serial transplantation 34. To test for self-renewal capacity, perform secondary transplantation. Sort human CD45+ cells from either the first primary recipients or enrich them using a mouse depletion kit following the manufacturer’s instructions. 35. Inject the recovered cells into a second recipient using the same protocol as described before (see steps 9 and 10). 36. After 6 to 12 weeks sacrifice the mice and analyze for human cells engraftment the same way as for primary transplantation (see steps 12 to 33). SUPPORT PROTOCOL 1

INTRA-BONE MARROW INJECTION To exclude stem cell homing interference and focus on the intrinsic capacity of a cell to self-renew, a few groups recently developed a highly sensitive strategy based on direct intra-bone marrow (IBM) injection of the candidate human stem cell (Mazurier et al., 2003; Wang et al., 2003; Yahata et al., 2003). IBM injection was found to be a more sensitive and adequate means to measure human HSC capacity. The intra-bone injection technique is performed under a short general anesthesia following a method originally described by Verlinden et al. (1998).

Additional Materials (see Basic Protocol) Anesthetic solution (see recipe) Post-operative analgesic (Vetergesic; Alstoe Animal health), diluted 1/10 in PBS and injected at 100 μl subcutaneously per mouse 29-G, 1/2 -in. needle (or 25-G needle) and insulin syringe (Tyco Healthcare) Anesthetize the mice 1. Inject the mice intraperitoneally with a dose of 0.2 to 0.25 ml of anesthetic solution. General anesthesia suppresses the heat-regulating mechanisms of the body. This is overcome by intra- and post-operative maintenance of body temperature in appropriate thermostatically controlled incubators or by other heat sources.

Xenotransplantation Model of Leukemia

2. Insert a syringe with a 25-G needle (maximum) into the joint surface of the right or left tibia/femur, and inject up to 40 μl cell suspension into the bone marrow cavity of the tibia or femur.

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3. During recovery, keep the animals under regular observation until full mobility is regained. 4. At this stage, provide at least one dose (100 μl) of post-operative analgesic (Vetergesic diluted 1/10) subcutaneously following bone marrow injection.

THAWING AML CELLS While freshly obtained AML cells are desirable for xenotransplantation, thawed frozen cells are also suitable, and it may be more convenient to collect and freeze the cells until one is ready for the transplantation experiment.

SUPPORT PROTOCOL 2

Materials AML cells, frozen in 1.8- to 2-ml cryovials DNase (Sigma, cat. no. D4513), thawed Fetal bovine serum (FBS; Stem Cell Technologies, cat. no. 06471) or any other suppliers Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) 37◦ C water bath 50-ml centrifuge tubes, sterile Table-top centrifuge equipped with swing-out bucket rotor for 15- and 50-ml conical tubes Cell strainer Additional reagents and equipment for counting cells (UNIT 1C.3) NOTE: Before thawing the samples, ensure the water bath is at 37◦ C. NOTE: Before starting, ensure that the DNase is completely thawed. 1. Rapidly thaw the AML cells (cryovial of 1.8- to 2-ml) in the 37◦ C water bath. There are no commercial suppliers of frozen AML. These samples can be obtained from clinics after informed consent from the patients has been obtained.

2. Add 100 ml DNase (1 mg/ml) dropwise into the cryovial. 3. Mix gently, wait 1 min, and transfer cells into a sterile 50-ml centrifuge tube. 4. Gently add 1 ml heat-inactivated, pure FBS dropwise, mix gently, and wait 1 min. 5. Slowly add 10 ml CMF-PBS/2% FBS and wait 1 min. 6. Slowly add up to 30 ml CMF-PBS/2% FBS to fill the tube. 7. Centrifuge 5 min at 200 × g, 4◦ C. 8. Resuspend in 1 ml CMF-PBS/2% FBS. 9. Filter using a cell strainer if needed (if cells are clumping). 10. Count viable cells (UNIT 1C.3) and use for purification or adoptive transfer protocols.

PURIFICATION STRATEGY Mononuclear cells from AML samples can be stained using a combination of antibodies. The most commonly used are the lineage cocktail antibodies (BD, cat. no. 340546), antiCD34 and anti-CD38. Stain the cells 25 to 30 min at 4◦ C in the presence of 5 μl/million of each of these antibodies. After the incubation period, centrifuge the cells 5 min at 380 × g, 4◦ C. Discard the supernatant, resuspend the cells in 1 ml CMF-PBS/2% FBS. Repeat this wash procedure one more time. After washing, the cells are ready to be sorted through a cell sorter (FACS Aria, BD, or equivalent).

SUPPORT PROTOCOL 3

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Anesthetic solution Mix 1 ml of Ketase solution (Fort Dodge Animal Health) with 0.5 ml of 2% Rompun solution (Bayer plc) and dilute with 8.5 ml of CMF-PBS. Store up to 1 month at 4◦ C. Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) Prepare 10× stock with 1.37 M NaCl, 27 mM KCl, 100 mM Na2 HPO2 , and 18 mM KH2 PO4 (adjust to pH 7.4 using HCl, if necessary) and autoclave. Store up to 2 months at room temperature. Prepare working solution by dilution of one part with nine parts water.

COMMENTARY Background Information

Xenotransplantation Model of Leukemia

AML-IC can be prospectively identified and purified as CD34+ /CD38− cells in AML patient samples, regardless of the phenotype of the bulk blast population, and these cells represent the only AML cells capable of self-renewal (Bonnet and Dick, 1997). The phenotype of AML-IC has been extended to include the expression of CD123 but the absence of CD71, HLA-DR, and CD117 (Blair et al., 1997, 1998; Blair and Sutherland, 2000; Jordan et al., 2000). In a recent study, this phenotype was further extended to include expression of CD33 and CD13 on AML-IC for the vast majority of patients (Taussig et al., 2005). Hence, the immunophenotype of the leukemic stem cell as defined by in vivo propagation is CD34+ /CD38− /CD71− / HLA-DR− /CD117− /CD33+ /CD13+ /CD123+ . However, the exclusivity of some of these markers is debatable. For instance, CD123 is indeed expressed on AML-IC, but it is also expressed on the vast majority of AML blasts (D. Bonnet, unpub. observ.) from most patients, and hence could be excluded from the above phenotype of AML-IC. Considerable heterogeneity within the AML-IC compartment exists. Lentiviral gene marking to track the behavior of individual LSCs, following serial transplantation, has revealed heterogeneity in their ability to selfrenew, similar to what is seen in the normal HSC compartment (Hope et al., 2004). Furthermore, using the NOD/SCID IL2Rgnull mice (see Anticipated Results) pretreated with anti-CD122, we show that in some patients the LSC activity can be detected in a CD34+ CD38+ population (Taussig et al., 2008). Thus, there is not a universal protocol

to purify for LSC in AML. Each AML patient sample should be tested first for its ability to engraft (prescreening) and secondly for identifying the nature of the cells responsible for repopulating ability. Secondary transplantation should also be used to test for the self-renewal ability of the LSC.

Critical Parameters By LSC, we refer to a cell that has selfrenewal and differentiation potential and is able to reinitiate the leukemia when transplanted into NOD/SCID mice. This definition does not preclude the nature of the cells being transformed. The confusion regarding the origin of the AML-IC may be due to the extreme heterogeneity of AML. Given the various possible routes to AML from a normal hematopoietic cell, it is not surprising that there is great heterogeneity in AML. Indeed, AML may be thought of as a large collection of different diseases that merely share similar characteristics. Indeed, the most effective risk stratification approach so far has been to examine the genetic abnormalities associated with a particular case of AML and compare it to previous experience with AML cases with the same abnormality (Grimwade et al., 1998, 2001). Although cytogenetic analysis allows the definition of the hierarchical groups with favorable, intermediate, and poor prognosis, the intermediate risk group contains patients with variable outcomes. Assessing the prognosis of this large group of patients is currently difficult.

Troubleshooting It is usually straightforward to detect AML engraftment. The human cells present express

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human CD45+ and the pan-myeloid marker CD33 without detection of lymphoid markers (CD19). Nevertheless, it happens that in some cases a chimeric engraftment can be detected, indicating that both human normal and leukemic cells are present in the engrafted mouse. In this case, it is usually important to confirm the leukemic origin of the CD33 subfraction by performing either FISH analysis or PCR for the mutations present in the original patient samples. Human CD45+ can be sorted based on CD19 and CD33, and both fractions should be tested for the presence of leukemic cells. Thus, it is not sufficient when screening for human AML engraftment to only test for the presence of human CD45+ cells, as in some cases these human cells could be exclusively normal cells.

Anticipated Results From AML patients’ samples at diagnosis, the capacity to engraft in the xenotransplantation model is usually ∼65% to 70%. Thus, there are still 25% to 30% of patients for which no engraftment could be detected after 10 to 12 weeks. The ability of a particular AML to engraft in the xenotransplantation model is related to the prognosis of individual AML cases (Pearce et al., 2006). Specifically, examination of the follow-up results from younger patients with intermediate-risk AML revealed a significant difference in overall survival between NOD/SCID-engrafting and non-engrafting cases. No differences have been detected between engrafting and nonengrafting cases in various engraftment variables including: homing ability, AML-IC frequency, and immune rejection by the host or alternative tissue sources. Hence, the ability to engraft NOD/SCID recipients seems to be an inherent property of the cells that is directly related to prognosis. Other mouse models have been developed to support the growth of human hematopoietic cells but less is known about the ability of these new models to sustain AML engraftment. The NOD/SCID-β2 microglobulin null (β2m−/− ) mouse has an additional defect in NK cell activity and is more tolerant of human grafts than the NOD/SCID model (Christianson et al., 1997; Kollet et al., 2000; Glimm et al., 2001). However, the percentage of AML samples that engraft in β2m−/− is similar to the level achieved using the NOD/SCID mice. Thus, it does not appear that the β2m−/− is superior for the engraftment of AML samples (Pearce et al., 2006). Furthermore, both NOD/SCID and β2m−/− are susceptible to developing lymphomas over time,

limiting their lifespan and preventing longterm reconstitution assessment. These hurdles have recently been overcome in three new strains: NOD/Shi-Scid IL2Rgnull (Yahata et al., 2002; Hiramatsu et al., 2003), NOD/SCID IL2Rgnull (Ishikawa et al., 2005; Shultz et al., 2005), and BALB/c-Rag2null IL2Rgnull (Traggiai et al., 2004), which all lack the IL-2 family common cytokine receptor gamma chain gene. The absence of functional receptors for IL2, IL-7, and other cytokines may prevent the expansion of NK cells and early lymphoma cells in NOD/SCID IL2Rgnull mice, resulting in better engraftment of transplanted human cells and longer lifespan of the mice. It was reported recently that human HSCs and progenitor cells engraft successfully in these mice and produce all human myeloid and lymphoid lineages. T and B cells migrate into lymphoid organs and mount HLA-dependent allogeneic responses, and generate antibodies against T cell–dependent antigens such as ovalbumin and tetanus toxin (Traggiai et al., 2004; Ishikawa et al., 2005). However, preliminary testing in our group seems to indicate that the NOD/SCID IL2Rgnull mice are not superior for leukemic engraftment to NOD/SCID or the NOD/SCID-β2m−/− mice. Thus, intrinsic properties of AML cells will dictate whether or not the cells will engraft.

Time Considerations It usually takes 4 to 6 weeks to detect engraftment. Nevertheless, we have seen in the past that the kinetics of AML engraftment, in contrast to normal stem cells, is usually slower, and thus it is better to wait 10 to 12 weeks for estimating the engraftment of AML samples. It sometimes happens, especially with poor cytogenetic samples that the mice get sick after only 3 to 4 weeks due to an infiltration of AML cells into solid organs (like spleen, liver, kidney). In this case, the mice have to be sacrificed earlier.

Acknowledgments The author would like to thank Dr. Daniel Pearce for his assistance in the preparation of this manuscript.

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in vivo have the phenotype CD34(+)/CD71 (-)/HLA-DR. Blood 92:4325-4335. Blair, A. and Sutherland, H.J. 2000. Primitive acute myeloid leukemia cells with long-term proliferative ability in vitro and in vivo lack surface expression of c-kit (CD117). Exp. Hematol. 28:660-671. Bonnet, D. and Dick, J.E. 1997. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat. Med. 3:730-737. Christianson, S.W., Greiner, D.L., Hesselton, R.A., Leif, J.H., Wagar, E.J., Schweitzer, I.B., Raian, T.V., Gott, B., Roopenian, D.C., and Shultz, L.D. 1997. Enhanced human CD4+ T cell engraftment in beta2-microglobulin-deficient NODscid mice. J. Immunol. 158:3578-3586. Donovan, J. and Brown, P. 2006a. Parenteral injections. Curr. Protoc. Immunol. 73:1.6.1-1.6.10. Donovan, J. and Brown, P. 2006b. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Glimm, H., Eisterer, W., Lee, K., Cashman, J., Holyoake, T. L., Nicolini, F., Shultz, L.D., von Kalle, C., and Eaves, C.J. 2001. Previously undetected human hematopoietic cell populations with short-term repopulating activity selectively engraft NOD/SCID-beta2 microglobulinnull mice. J. Clin. Invest. 107:199-206. Grimwade, D., Walker, H., Oliver, F., Wheatley, K., Harrison, C., Harrison, G., Rees, J., Hahn, I., Stevens, R., Burnett, A., and Gladstone, A. 1998. The importance of diagnostic cytogenetics on outcome in AML: Analysis of 1,612 patients entered into the MRC AML 10 trial. The Medical Research Council Adult and Children’s Leukaemia Working Parties. Blood 92:23222333. Grimwade, D., Walker, H., Harrison, G., Oliver, F., Chatters, S., Harrison, C. J., Wheatley, K., Burnett, A.K., and Goldstone, A.H.; Medical Research Council Adult Leukemia Working Party. 2001. The predictive value of hierarchical cytogenetic classification in older adults with acute myeloid leukemia (AML): Analysis of 1065 patients entered into the United Kingdom Medical Research Council AML11 trial. Blood 98:1312-1320. Hiramatsu, H., Nishikomori, R., Heike, T., Ito, M., Kobayashi, K., Katamura, K., and Nakahata, T. 2003. Complete reconstitution of human lymphocytes from cord blood CD34+ cells using the NOD/SCID/gammacnull mice model. Blood 102:873-880. Hope, K. J., Jin, L., and Dick, J. E. 2004. Acute myeloid leukemia originates from a hierarchy of leukemic stem cell classes that differ in self-renewal capacity. Nat. Immunol. 5:738743.

Xenotransplantation Model of Leukemia

Ishikawa, F., Yasukawa, M., Lyons, B., Yoshida, S., Miyamoto, T., Yoshimoto, G., Watanabe, T., Akashi, K., Shultz, L.D., and Harada, M. 2005. Development of functional human blood and immune systems in NOD/SCID/IL2 receptor {gamma} chain(null) mice. Blood 106:15651573.

Jordan, C.T., Upchurch, D., Szilvassy, S.J., Guzman, M.L., Howard, D.S., Pettigrew, A.L., Meyerrose, T., Rossi, R., Grimes, B., Rizzeri, D.A., Luger, S.M., and Phillips, G.L. 2000. The interleukin-3 receptor alpha chain is a unique marker for human acute myelogenous leukemia stem cells. Leukemia 14:1777-1784. Lapidot, T., Sirard, C., Vormoor, J., Murdoch, B., Hoang, T., Caceres-Cortes, J., Minden, M., Paterson, B., Caligiuri, M.A., and Dick, J.E. 1994. A cell initiating human acute myeloid leukaemia after transplantation into SCID mice. Nature 367:645-648. Kollet, O., Peled, A., Byk, T., Ben-Hur, H., Greiner, D., Shultz, L., and Lapidot, T. 2000. beta2 microglobulin-deficient (B2m(null)) NOD/SCID mice are excellent recipients for studying human stem cell function. Blood 95:3102-3105. Mazurier, F., Doedens, M., Gan, O.I., and Dick, J.E. 2003. Rapid myeloerythroid repopulation after intrafemoral transplantation of NOD-SCID mice reveals a new class of human stem cells. Nat. Med. 9:959-963. Pearce, D.J., Taussig, D., Zibara, K., Smith, L.L., Ridler, C.M., Preudhomme, C., Young, B.D., Rohatneer, A.Z., Lister, T.A., and Bonnet, D. 2006. AML engraftment in the NOD/SCID assay reflects the outcome of AML: Implications for our understanding of the heterogeneity of AML. Blood 107:1166-1173. Shultz, L.D., Lyons, B.L., Burzenski, L.M., Gott, B., Chen, X., Chaleff, S., Kotb, M., Gillies, S.D., King, M., Mangada, J., Greiner, D.L., and Handgretinger, R. 2005. Human lymphoid and myeloid cell development in NOD/LtSz-scid IL2R gamma null mice engrafted with mobilized human hematopoietic stem cells. J. Immunol. 174:6477-6489. Taussig, D.C., Pearce, D.J., Simpson, C., Rohatiner, A.Z., Lister, T.A., Kelly, G., Luongo, J.L., Danet-Desnoyers, G.A., and Bonnet, D. 2005. Hematopoietic stem cells express multiple myeloid markers: Implications for the origin and targeted therapy of acute myeloid leukemia. Blood 106:4086-4092. Taussig, D.C., Miraki-Moud, F., Anjos-Afonso, F., Pearce, D.J., Allen, K., Ridler, C., Lillington, D., Oakervee, H., Cavenagh, J., Agrawal, S.G., Lister, T.A., Gribben, J.G., and Bonnet, D. 2008. Anti-CD38 antibody-mediated clearance of human repopulating cells masks the heterogeneity of leukemia-initiating cells. Blood 112:568575. Traggiai, E., Chicha, L., Mazzucchelli, L., Bronz, L., Piffaretti, J.C., Lanzavecchia, A., and Manz, M.G. 2004. Development of a human adaptive immune system in cord blood cell-transplanted mice. Science 304:104-107. Verlinden, S.F., van Es, H.H., and van Bekkum, D.W. 1998. Serial bone marrow sampling for long-term follow up of human hematopoiesis in NOD/SCID mice. Exp. Hematol. 26:627-630. Wang, J., Kimura, T., Asada, R., Harada, S., Yokota, S., Kawamoto, Y., Fijimura, Y., Tsuji,

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T., Ikehara, S., and Sonoda, Y. 2003. SCIDrepopulating cell activity of human cord bloodderived CD34- cells assured by intra-bone marrow injection. Blood 101:2924-2931. Yahata, T., Ando, K., Nakamura, Y., Ueyama, Y., Shimamura, K., Tamaoki, N., Kato, S., and Hotta, T. 2002. Functional human T lymphocyte development from cord blood CD34+ cells in nonobese diabetic/Shi-scid, IL-2 receptor gamma null mice. J. Immunol. 169:204-209. Yahata, T., Ando, K., Sato, T., Miyatake, H., Nakamura, Y., Muguruma, Y., Kato, S., and Hotta, T. 2003. A highly sensitive strategy for SCID-repopulating cell assay by direct injection of primitive human hematopoietic cells into NOD/SCID mice bone marrow. Blood 101:2905-2913.

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Culture and Isolation of Brain Tumor Initiating Cells

UNIT 3.3

Monika Lenkiewicz,1 Na Li,1 and Sheila K. Singh1 1

McMaster University, Hamilton, Ontario, Canada

ABSTRACT This unit describes protocols for the culture and isolation of brain tumor initiating cells (BTIC). The cancer stem cell (CSC) hypothesis suggests that tumors are maintained exclusively by a rare fraction of cells that have stem cell properties. We applied culture conditions and assays originally used for normal neural stem cells (NSCs) in vitro to a variety of brain tumors. The BTIC were isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133. Only the CD133+ brain tumor fraction contains cells capable of sphere formation and sustained self-renewal in vitro, and tumor initiation in NOD-SCID mouse brains. Therefore, CD133+ BTICs satisfy the definition of cancer stem cells in that they are able to generate a replica of the patient’s tumor and they exhibit self-renewal ability through serial retransplantation. This established that only a rare subset of brain tumor cells with stem cell properties are tumor-initiating, and, in this unit, we describe their culture and isolation. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 11:3.3.1-3.3.10.  Keywords: brain tumor initiating cells (BTICs) r tumor sphere culture r CD133 r cell sorting r cancer stem cell (CSC)

INTRODUCTION In this unit, protocols for the culture and isolation of brain tumor initiating cells (BTICs) are described. The cancer stem cell (CSC) hypothesis suggests that tumors are maintained exclusively by a rare fraction of cells that have stem cell properties. Here, we discuss the methods that we first used to prospectively identify and enrich for a subpopulation of human BTICs that exhibit the stem cell properties of proliferation, self-renewal, and differentiation in vitro (Singh et al., 2003) and in vivo (Singh et al., 2004). We applied culture conditions and assays originally used to characterize normal neural stem cells (NSCs) in vitro (Reynolds and Weiss, 1992; Tropepe et al., 1999) to a variety of pediatric and adult brain tumors. The BTIC were exclusively isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133 (Yin et al., 1997; Yu et al., 2002). Only the CD133+ brain tumor fraction contains cells that are capable of sphere formation and sustained self-renewal in vitro, as well as tumor initiation in NOD-SCID mouse brains. Therefore, CD133+ BTICs satisfy the definition of cancer stem cells in that they are able to generate a replica of the patient’s tumor and they exhibit self-renewal ability both in vitro and in vivo through serial retransplantation (Bonnet and Dick, 1997; Reya et al., 2001). This formally established that only a rare subset of brain tumor cells with stem cell properties are tumor-initiating. This unit begins with a method for the culture of tumor spheres from human brain tumors (Basic Protocol 1) and follows with a protocol for the prospective isolation of BTICs from these cultures by magnetic bead sorting (Basic Protocol 2) or fluorescence activated sorting (Alternate Protocol) for CD133. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. Current Protocols in Stem Cell Biology 3.3.1-3.3.10 Published online October 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0303s11 C 2009 John Wiley & Sons, Inc. Copyright 

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NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

CULTURE OF TUMOR SPHERES FROM HUMAN BRAIN TUMORS This protocol is adapted from those previously established for isolation of neural stem cells as neurospheres (Reynolds and Weiss, 1992, 1996; Reynolds et al., 1992), and has been applied to the culture of human brain tumors. We use this culture method specifically to select for cell populations within human brain tumors that possess stem cell properties. Serum-free medium (SFM) allows for the maintenance of an undifferentiated stem cell state, and the addition of bFGF and EGF allows for the proliferation of multipotent, selfrenewing, and expandable tumor spheres. The medium on these tumor sphere cultures should be changed every other day, and when primary tumor spheres reach a critical size of >100 μm, they may be passaged to secondary spheres based on the principles of the neurosphere assay. The frequency of the stem cell population within the brain tumor can be determined by primary sphere formation assay, and the minimal frequency of repopulating tumor sphere cells within the culture can be estimated by serial sphere formation through limiting dilution analysis (Tropepe et al., 1999).

Materials Hi/low aCSF (see recipe) 95% O2 /5% CO2 Enzyme digestive mix for tumors (see recipe) or high-performance liquid chromatography–purified collagenase/dispase cocktail (e.g., Liberase Blendzyme 3 from Roche) Human brain tumor specimen Tumor sphere medium (see recipe) Soybean trypsin inhibitor (from Glycine max; Sigma, cat. no. T9003) 10 ng/μl leukemia inhibitor factor (LIF) stock solution (see recipe) 10 ng/μl recombinant human basic fibroblast (bFGF) stock solution (see recipe) 10 ng/μl recombinant human epidermal growth factor (EGF) stock solution (see recipe) 0.22-μm pore size, 150-ml filter system (Corning) Oxygen tank setup in laminar flow hood to allow for sterile oxygenation of solutions 15- and 50-ml conical centrifuge tubes 0.22-μm syringe filter units (Millipore) 10-cm2 tissue culture–grade plates (Falcon) or 100-mm Ultra Low Attachment Culture dishes (Corning) Fine sterile scissors and forceps Incubator-shaker (VWR Scientific) Tabletop centrifuge 70-μm cell strainer (BD Falcon) Dissociate human brain tumor tissue 1. Prepare 125 ml of hi/low aCSF as described in Reagents and Solutions, and filter sterilize using a 0.2-μm filter and 150-ml filter system. Culture and Isolation of Brain Tumor Initiating Cells

2. Bubble aCSF with sterile 95% O2 /5% CO2 for 15 min and place in 37◦ C water bath.

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3. Freshly prepare the enzyme digestive mix for tumors (see Reagents and Solutions) by weighing out the enzymes (and kynurenic acid) into separate 15-ml centrifuge tubes. Resuspend each enzyme/reagent in 10-ml of hi/low aCSF, vortex thoroughly, and filter all of the components through a 0.22-μm syringe filter into one sterile 50-ml centrifuge tube. Alternatively, measure high performance liquid chromatography– purified collagenase/dispase cocktail (Liberase Blendzyme 3) according to the manufacturer’s instructions, and resuspend in aCSF. If using enzyme digestive mix for tumors, final working concentrations are: trypsin 1.33 mg/ml, hyaluronidase 0.67 mg/ml, and kynurenic acid 0.1 to 0.17 mg/ml. If using the alternate method with Liberase Blendzyme 3, the final working concentration is 0.2 W¨unsch units/ml in a total of 15 ml aCSF.

4. In sterile hood, wash tumor specimen in a 10-cm2 plate filled with hi/low aCSF, transferring sequentially to new 10-cm2 plates filled with aCSF until excess blood is thoroughly washed out. The authors have tested 12 brain tumor subtypes (Singh et al., 2003, 2004) and they all had some proportion of BTICs; sample size can be very small and BTICs are still isolated, so there is no minimum sample size. BTIC yield directly correlates with grade of tumor, i.e., there is lower BTIC yield with benign low-grade tumors and higher yield with biologically aggressive malignant tumors (Singh et al., 2003, 2004).

5. Using sterile fine scissors and forceps, cut tumor into 1-mm3 pieces in a 10-cm2 tissue culture plate with 2 to 3 ml of enzymatic digestion mixture (either the enzyme/ kynurenic acid mix used in step 3, or Liberase Blendzyme 3). 6. Collect tumor pieces with 10-ml pipet, pipetting up and down and dispensing the tumor fragments into 30 ml of the enzymatic digestion mixture used in step 3 or 15 ml Liberase Blendzyme 3. 7. Digest at 37◦ C for 30 to 90 min (depending on tumor size) with gentle mixing in an incubator-shaker. The incubator-shaker provides superior digestion and subsequent yield of cells compared to a rocker. If using Liberase Blendzyme 3, the digestion period can be reduced to 15 to 30 min.

Stop enzymatic reaction Steps 8 to 10 are skipped if using Liberase Blendzyme 3. In that case, simply filter the cell digest from step 7 through a 70-μm cell strainer and proceed to step 11. 8. Freshly prepare the trypsin inhibitor solution by resuspending 35 mg of trypsin inhibitor in 5 ml tumor sphere medium (without LIF, bFGF, or EGF) and filtering through a 0.2-μm filter. 9. Centrifuge cells 3 min at ∼450 × g, room temperature, and take off as much supernatant as possible without dislodging the tumor tissue at the bottom of the tube. 10. Add the 5 ml of trypsin inhibitor solution from step 8, mix well, and filter through a 70-μm cell strainer.

Plate cells in tumor sphere medium with growth factors 11. Centrifuge cells 3 min at ∼450 × g, room temperature. Aspirate all of the supernatant and resuspend in 10 to 12 ml tumor sphere medium supplemented with 10 ng/ml LIF (add from 10 ng/μl stock), 20 ng/ml bFGF (add from 10 ng/μl stock), and 20 ng/ml EGF (add from 10 ng/μl stock). 12. Plate cells in 10-cm2 dishes at 2 × 105 cells per cm2 , in 10 to 12 ml of tumor sphere medium with LIF, bFGF, and EGF. Incubate in a 37◦ C 5% CO2 incubator.

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A 100x

400x

0.05 mm

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200x

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Figure 3.3.1 Brain tumor initiating cells forming sphere-like structures in vitro. (A) Tumor spheres of anaplastic medulloblastoma. (B) Tumor spheres of metastatic melanoma.

13. Feed cells every other day by centrifuging 3 min at 450 × g, room temperature, aspirating the medium, and replacing it with fresh tumor sphere medium supplemented with fresh LIF, bFGF, and EGF, as described in step 11. 14. When number of spheres per plate has doubled or spheres are consistently >120 μm in size, split cultures by repeating steps 11 to 12. Figure 3.3.1 shows brain tumor initiating cells forming sphere-like structures in vitro.

BASIC PROTOCOL 2

Culture and Isolation of Brain Tumor Initiating Cells

ENRICHMENT OF BTICs BY MAGNETIC BEAD SORTING FOR CD133 This protocol is used for the prospective isolation or enrichment of BTICs from tumor sphere cultures, which constitute primary human brain tumor cells, cultured as per Basic Protocol 1. Cell sorting is performed as soon as tumor spheres begin to form in culture, and is optimally performed within 1 to 24 hr after initial cell culture. Cells must be in single-cell suspension for optimal sorting, and spheres are gently triturated or chemically dissociated prior to cell sorting, by methods detailed below. Cell sorting for CD133 can be performed either by magnetic bead cell sorting (MBCS; this protocol) or by fluorescence activated cell sorting (FACS; Alternate Protocol), and we provide methods for each of these options. Considerations of experimental timing, cell sorter availability at short notice or at night, cell viability after processing through FACS, individual tumor characteristics, overall cell number, and inter-user reliability can help to discern which method of cell sorting should be used for each tumor sphere sample.

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Materials Primary human brain tumor cells growing in culture (Basic Protocol 1) Incubation buffer: phosphate buffered saline (PBS; Invitrogen, cat. no. 14190-144)/0.5% (w/v) BSA with (or without; see below) 2 mM disodium EDTA, pH 7.2 FcR blocking reagent (Miltenyi Biotec) Phosphate-buffered saline (Invitrogen, cat. no. 14190-144) MACS CD133 Cell Isolation Kit (Miltenyi Biotec) consisting of: beads conjugated to monoclonal mouse anti–human CD133 Microbeads, Isotype IgG1 magnetic cell separator (MiniMACS column magnet) MS separation columns CD133-2-PE antibody mouse IgG2b-PE isotype control antibody Tumor sphere medium (see recipe) 4% (w/v) paraformaldehyde (optional) 15-ml conical centrifuge tubes (Falcon or equivalent) Flame-narrowed pipets 70-μm cell strainer 6-well Ultra Low Cluster plates (Corning) Flow cytometry tubes (BD Falcon 352058) Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) and flow cytometry (Robinson et al., 2009) NOTE: Work quickly and keep cells/buffer (not culture media) cold (4◦ to 6◦ C).

Prepare cell suspensions 1. Place 2 × 105 cultured tumor spheres in a 15-ml centrifuge tube. Centrifuge cells 3 min at ∼450 × g, room temperature, and remove medium. Resuspend cell pellet in 1 ml of incubation buffer. Do not use EDTA if working with Notch pathway molecules, due to potential interactions. RBC contamination will decrease purity and cause tumor cell death upon RBC lysis; to remove RBCs if specimen is vascular, treat the cells with an RBC lysis buffer (StemCell Technologies, cat. no. 67850).

2. Triturate gently with flame-narrowed pipet or micropipet tip (if not at single-cell suspension already). 3. Filter through 70-μm cell strainer, and count viable cells using a hemacytometer and trypan blue (UNIT 1C.3). Gently triturate and strain cells just before flow to avoid clumping. You may use a flamenarrowed or regular pipet or micropipet; we avoid excessive trituration to minimize trauma to the cells.

4. Aliquot equal amounts of cell suspension into four microcentrifuge tubes as follows:

negative control (unstained) (A1) isotype control (A2) pre-sort CD133-1 staining of specimen (A3) bulk of the cells in the last aliquot, for bead sorting (A4). Perform magnetic separation 5. Resuspend A1 and A4 in 300 μl incubation buffer and add 100 μl FcR blocking reagent to A4. If cell number in A4 is >5 × 106 , divide into two tubes and treat both the same way. Resuspend A2 and A3 in 500 μl PBS. Keep aliquots A1, A2, and A3 on ice. Current Protocols in Stem Cell Biology

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6. Add 1 μl beads per 106 cells to tube A4. Mix beads well in A4 and leave at 4◦ C (in the refrigerator in dark) for 30 min. 7. After 30 min, take the A4 aliquot and place the microcentrifuge tube into a MACS separator magnet. Use LS columns or MS columns provided with the kit, based on cell number. 8. Rinse column with 3 ml incubation buffer for LS column. Add 3 ml cell suspension onto the column, and allow negative cells to pass through. Collect negative cells in a 15-ml centrifuge tube. 9. Wash four times, each time with 3 ml incubation buffer. Pool with the negative cells in the 15-ml centrifuge tube from step 8. 10. Remove column from separator and place on a clean 15-ml centrifuge tube. Flush out positive fraction with 3 ml incubation buffer by firm application of hand to column. 11. Repeat, and apply column one more time to resuspended CD133+ cells, to purify this population further if cell numbers permit. Reapplying the cells to the column increases the purity of the CD133+ cells to ∼95%.

12. Centrifuge the sorted A4 aliquots, CD133− from steps 8 and 9 and CD133+ from step 11, 3 min at 450 × g, room temperature, and remove the supernatant. Wash by adding 2 ml incubation buffer, centrifuging again as before, and removing the supernatant. Resuspend each pellet in 3 ml tumor sphere medium. 13. Plate 2–5 × 104 cells per well in 3 ml tumor sphere medium per well of a 6-well plate. Re-equilibrate immediately to 37◦ C.

Perform a purity check by immunostaining and flow cytometric analysis 14. Take a 10-μl aliquot from the last CD133+ and CD133− aliquots (step 12) and resuspend each in 500 μl PBS. 15. Add 5 μl CD133-2-PE antibody to each sample, and incubate at 4◦ C for 30 min. These specimens will be taken to the flow cytometry lab for a purity check of the CD133+ and CD133− sorted populations.

16. Add 5 to 10 μl of CD133-2-PE antibody to tube A2 and 5 to 10 μl isotype IgG2b-PE control antibody to tube A3 (see step 4). Incubate at room temperature for 15 to 30 min. Also incubate the unstained A1 control and carry it through the remaining steps. 17. Centrifuge aliquots A2 and A3 3 min at ∼450 × g, room temperature, and remove the supernatants. Wash by adding 10 to 20 pellet volumes of PBS, centrifuging again as before, and removing the supernatant. Finally, resuspend in 250 μl PBS. The cells should be washed as thoroughly as possible after staining; however, this must be balanced with the potential for cell loss.

18. Optional: Post-fix A2, A3, and A1 aliquots and the two purity check aliquots from A4 in 2% (w/v) paraformaldehyde (i.e., add 250 μl 4% paraformaldehyde to each aliquot). 19. Triturate each sample gently and transfer to flow cytometry tubes, preferably through strainer caps to remove clumps. Culture and Isolation of Brain Tumor Initiating Cells

20. Analyze the five samples by flow cytometry (Robinson et al., 2009). See Figure 3.3.2 for sample results.

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A tumor cells

negative control (A1) IgG2b isotype control (A2) CD133-2-PE (A3)

magnetic sorting (A4) (Miltenyi CD 133 isolation kit) CD133 cells

CD133 cells

purity check (flow cytometry)

B 64

64 95% ( )

Events

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0 100

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0 104 100

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Figure 3.3.2 (A) Histogram of magnetic bead separation protocol for CD133. (B) Histograms of flow cytometry for the unsorted tumor cells (left) and sorted CD133+ cells (right) from a 49-year-old female metastatic melanoma patient. Corresponding isotype controls were overlaid as the unshaded histograms.

ENRICHMENT OF BTICs BY FLUORESCENCE ACTIVATED CELL SORTING

ALTERNATE PROTOCOL

Cell preparation and staining are performed as per the magnetic separation protocol (Basic Protocol 2), with the exception that the A4 aliquot, containing the bulk of the cells, is used for fluorescence activated cell sorting (FACS; Robinson et al., 2009). This aliquot should be resuspended in 300 μl PBS with 100 μl FcR blocking reagent. If the cell number in A4 is >5 × 106 , divide the suspension into two tubes and treat both the same way. Incubate A4 with a 1:100 dilution of CD133-2-PE antibody and leave at 4◦ C (in refrigerator in dark) for 30 min. Wash and centrifuge in 10 to 20 volumes of PBS as described in Basic Protocol 2, and resuspend A4 cells in 250 μl PBS. Take A4 aliquot to your FACS operator, with aliquots A1 through A3 prepared as in Basic Protocol 2 for controls.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Enzyme digestive mix for tumors 40 mg bovine pancreas trypsin (Sigma, cat no. T9201) 20 mg bovine testis hyaluronidase (Sigma, cat no. H3884) 3 to 5 mg kynurenic acid (Sigma, cat no. K3375) Current Protocols in Stem Cell Biology

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Resuspend these three components in 30 ml of warm (37◦ C) hi/low aCSF and filter through a 0.2-μm filter. This enzyme digestive mix must be made up fresh prior to use.

Hi/low artificial cerebral spinal fluid (aCSF) For a total of 125 ml aCSF, combine the following: 7.75 ml 2 M NaCl 0.625 ml 1 M KCl 0.4 ml 1 M MgCl2 21.125 ml 155 mM NaHCO3 1.25 ml 1 M glucose 0.1157 ml 108 mM CaCl2 93.73 ml H2 O Hi/low aCSF can be prepared and stored at 4◦ C for up to 6 months (Dr. L. Doering, pers. comm.).

Recombinant human basic fibroblast growth factor (bFGF) stock solution, 10 ng/μl Resuspend lyophilized bFGF (Invitrogen) to a final concentration of 10 ng/μl in PBS (Invitrogen, cat. no. 14190-144) containing 0.1% (w/v) BSA. Store at −30◦ C.

Recombinant human epidermal growth factor (EGF) stock solution, 10 ng/μl Resuspend lyophilized EGF (Sigma) to a final concentration of 10 ng/μl in PBS (Invitrogen, cat. no. 14190-144) containing 0.1% (w/v) BSA. Store at −30◦ C.

Tumor sphere medium DMEM/F12 (Invitrogen) containing: 1× antibiotic-antimycotic (Wisent, cat. no. 450-115-EL; http://www.wisent.ca/) 1× hormone mix (N2 Supplement; Invitrogen, cat. no. 17502-048) 10 mM HEPES (Wisent, cat no. 330-050; http://www.wisent.ca/) 0.6% (w/v) glucose 60 μg/ml N-acetylcysteine (Sigma, cat no. A9165) 2% (w/v) NSF-1 (neural cell survival factor; Lonza, cat. no. CC-4323) Store for up to several weeks at 4◦ C Supplement the medium just before use with: 10 ng/ml LIF 20 ng/ml bFGF (see recipe for stock solution) 20 ng/ml EGF (see recipe for stock solution) COMMENTARY Background Information

Culture and Isolation of Brain Tumor Initiating Cells

When multipotent NSCs were isolated from the mammalian neuraxis more than a decade ago, culture conditions were developed that allowed embryonic EGF-responsive cells to proliferate as floating spheres (neurospheres), which could be easily manipulated for subsequent passage and differentiation (Reynolds and Weiss, 1992; Reynolds et al., 1992). Serum-free medium (SFM) allowed for the maintenance of an undifferentiated state, and the addition of saturating concentrations of bFGF and EGF (20 ng/ml) induced the proliferation of multipotent, self-renewing and ex-

pandable neural stem cells (Reynolds et al., 1992; Reynolds and Weiss, 1996). This neurosphere culture system and analysis procedure to identify NSCs has permitted in vitro characterization of these cells, but in a retrospective fashion, as the multipotential floating clusters of cells are inferred to have been derived from clonal expansion of a single NSC (Tropepe et al., 1999). Prospective study of this cell has been previously limited by lack of cell surface markers necessary for its isolation; recent reports show NSC enrichment using antibodies against the cell surface protein CD133 (Yin et al., 1997; Yu et al., 2002). Uchida and

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colleagues determined that this 120-kDa fivetransmembrane cell surface receptor of unknown function could effectively sort sphereforming cells from their non-sphere-forming counterparts in isolates of fetal human brain. Normal CD133+ human fetal brain cells not only efficiently form neurospheres in vitro, but also demonstrate the key stem cell properties of self-renewal and multilineage differentiation, and are capable of seamless lifelong engraftment and multilineage contribution to the mouse brain (Uchida et al., 2000). These findings represented the first evidence that the in vitro neurosphere-forming cell, when prospectively isolated, bore key stem cell properties both in vitro and in vivo. We applied the neurosphere culture assay to human brain tumors of different phenotypes, in order to select for stem cell growth and functionally characterize human brain tumor cell populations. Regardless of pathological subtype, within 24 to 48 hr of primary culture most brain tumors yielded a minority fraction of cells that demonstrated growth into clonally derived neurosphere-like clusters, termed tumor spheres. Tumor spheres are defined as clonally derived nonadherent colonies of cells derived from a single tumor stem cell. The remaining majority of tumor cells exhibited adherence, loss of proliferation, and subsequent differentiation, whereas tumor spheres remained nonadherent, continuing exclusively to self-renew and expand the tumor cell culture. From these cultures, the BTIC can then be exclusively isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133 (Yin et al., 1997; Yu et al., 2002). Only the CD133+ brain tumor fraction contains cells that are capable of sphere formation and sustained self-renewal in vitro, and tumor initiation in NOD-SCID mouse brains (Singh et al., 2003, 2004). Therefore, our characterization of CD133+ BTICs lends support for the application of the cancer stem cell hypothesis to solid tumors, and our in vitro and in vivo BTIC models will provide the foundation for a brain cell hierarchy that may begin to organize the heterogeneity of brain tumors.

Critical Parameters and Troubleshooting There are many different methods of culturing brain tumor cells, and many variable applications of the neurosphere assay to human brain tumors. In culturing cells with stem cell properties in both normal and cancer tissues, there is no standardized protocol with respect

to growth factors, hormones, and their concentrations (Chaichana et al., 2006). In establishing human tumor sphere cultures, we have found that our tumor sphere medium and its components have been optimized for growth of healthy spheres with good cell viability and reliable stem cell frequency across different tumor subtypes. We recommend plating the cells at a high density (e.g., 2 × 105 cells per cm2 ), and we anticipate a large amount of cell death in the first few days of culture. This cell death can be attributed to both the elevated apoptotic activity of cancer cells and to the fact that the bulk tumor population will not survive in serum-free conditions, which does select for stem and progenitor cell growth. Another parameter within the tumor culture protocol that requires much testing and optimization is the enzyme digestion. Both the length of time for tumor digestion and the choice of enzyme/protease mix will influence the yield of tumor cells and their viability. Length of time for tumor specimen digestion must be judged based on each individual tumor, with larger and firmer specimens requiring longer digestion. Tumor pathological subtype may also influence this decision, as some benign brain tumors have a more extensive collagen- or fibrin-based framework, and some malignant tumors may harbor a greater degree of pre-existing tissue necrosis. Underdigestion will not yield the largest possible cell number, whereas overdigestion results in DNA lysis and increased amounts of stringy white fibers and cellular debris in the culture. In terms of choice of enzyme/protease mix for digestion, we have begun to favor the use of collagenase-based cocktails, due to the fact that CD133 is trypsin-sensitive (Fukuchi et al., 2004; Schwab et al., 2008), and this receptor may be cleaved during the digestion process. If the cells are not stringently treated with antitrypsin or are not allowed to re-equilibrate for a long enough time in tumor sphere medium post-digestion, the CD133 expression level on cell sorting may be underestimated. For sorting either with FACS or MBCS, it is critical to obtain a single-cell suspension, and neither normal neurospheres nor adherent tumor spheres are easily amenable to dissociation. Thus, we use a combination of very gentle mechanical trituration just prior to sorting, with filtering through cell-strainer caps into flow cytometry tubes, to prevent clumping during the sort. We also recommend treating the cells with chemical cell dissociation buffers, which may provide a more gentle alternative to mechanical dissociation.

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Anticipated Results Depending on the size of the original tumor specimen, our tissue culture protocol should yield 1–100 × 106 viable tumor sphere cells from brain tumors of different subtypes, which then can be subjected to a panel of in vitro stem cell assays, including primary sphere formation assays, proliferation assays, limitingdilution assays, and differentiation assays. Our BTIC enrichment protocol should provide millions of CD133− brain tumor non-stem cells, and thousands to millions of CD133+ BTICs, depending on the clonogenic frequency and CD133 index of each tumor subtype. In general, the increased self-renewal capacity of the BTIC and the correlated CD133 index are highest from the most aggressive clinical samples of medulloblastoma and glioblastoma compared with low-grade gliomas. Both bead and FACS separation routinely yield average purities of 80% to 90% for the CD133+ cells and >99.5% purity for CD133− cells; thus, these methods should be considered as good enrichment methods, not methods for isolation to purity of BTICs.

Time Considerations Tumor dissociation and plating of cells into tumor sphere cultures typically takes 2 to 4 hr, and cell sorting can be completed within 4 hr.

Literature Cited Bonnet, D. and Dick, J.E. 1997. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat. Med. 3:730-737. Chaichana, K., Zamora-Berridi, G., CamaraQuintana, J., and Qui˜nones-Hinojosa, A. 2006. Neurosphere assays: Growth factors and hormone differences in tumor and nontumor studies. Stem Cells 24:2851-2857. Fukuchi, Y., Nakajima, H., Sugiyama, D., Hirose, I., Kitamura, T., and Tsuji, K. 2004. Potential human placenta-derived cells have mesenchymal stem/progenitor cell. Stem Cells 22:649-658. Reya, T., Morrison, S.J., Clarke, M.F., and Weissman, I.L. 2001. Stem cells, cancer, and cancer stem cells. Nature 414:105-111.

Reynolds, B.A. and Weiss, S. 1992. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707-1710. Reynolds, B.A., Tetzlaff, W., and Weiss, S.A. 1992. Multipotent EGF-responsive striatal embryonic progenitor cell produces neurons and astrocytes. J. Neurosci. 12:4565-4574. Reynolds, B.A. and Weiss, S. 1996. Clonal and population analyses demonstrate that an EGFresponsive mammalian embryonic CNS precursor is a stem cell. Dev. Biol. 175:1-13. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. 2009. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Schwab, K.E., Hutchinson, P., and Gargett, C.E. 2008. Identification of surface markers for prospective isolation of human endometrial stromal colony-forming cells. Hum. Reprod. 23:934-943. Singh, S.K., Clarke, I.D., Terasaki, M., Bonn, V.E., Hawkins, C., Squire, J., and Dirks, P.B. 2003. Identification of a cancer stem cell in human brain tumors. Cancer Res. 63:5821-5828. Singh, S.K., Hawkins, C., Clarke, I.D., Squire, J.A., Bayani, J., Hide, T., Henkelman, R.M., Cusimano, M.D., and Dirks, P.B. 2004. Identification of human brain tumor initiating cells. Nature 432:396-401. Tropepe, V., Sibilia, M., Ciruna, B.G., Rossant, J., Wagner, E.F., and van der Kooy, D. 1999. Distinct neural stem cells proliferate in response to EGF and FGF in the developing mouse telencephalon. Dev. Biol. 208:166-188. Uchida, N., Buck, D.W., He, D., Reitsma, M.J., Masek, M., Phan, T.V., Tsukamoto, A.S., Gage, F.H., and Weissman, I.L. 2000. Direct isolation of human central nervous system stem cells. Proc. Natl. Acad. Sci. U.S.A. 97:14720-14725. Yin, A.H., Miraglia, S., Zanjani, E.D., AlmeidaPorada, G., Ogawa, M., Leary, A.G., Olweus, J., Kearney, J., and Buck, D.W. 1997. AC133, a novel marker for human hematopoietic stem and progenitor cells. Blood 90:5002-5012. Yu, Y., Flint, A., Dvorin, E.L., and Bischoff, J. 2002. AC133-2, a novel isoform of human AC133 stem cell antigen. J. Biol. Chem. 277:2071120716.

Culture and Isolation of Brain Tumor Initiating Cells

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Human iPS Cell Derivation/Reprogramming

UNIT 4A.1

In-Hyun Park1 and George Q. Daley1, 2 1

Children’s Hospital Boston and Dana-Farber Cancer Institute, Harvard Medical School, Harvard Stem Cell Institute, Boston, Massachusetts 2 Brigham and Women’s Hospital, Howard Hughes Medical Institute, Boston, Massachusetts

ABSTRACT This unit describes a protocol for deriving induced pluripotent stem (iPS) cells from human fibroblast cells. Human fibroblast cells are infected with retroviral vectors expressing four transcription factors (Oct4, Sox2, Klf4, and Myc) and selected for 3 to 4 weeks under human embryonic stem (hES) cell culture conditions. iPS cell colonies are mechanically isolated using a dissection microscope and handled like hES cells thereafter. Human iPS cells share similarities with hES cells including the expression of pluripotency genes, and differentiation as embryoid bodies in vitro into three germ layers (EB) and in vivo C 2009 by John Wiley & as teratomas. Curr. Protoc. Stem Cell Biol. 8:4A.1.1-4A.1.8.  Sons, Inc. Keywords: human induced pluripotent stem (iPS) cells r human embryonic stem (hES) cells r reprogramming r retroviral vectors

INTRODUCTION This unit describes a protocol for deriving induced pluripotent stem (iPS) cells from human fibroblasts. hES cells have the property of self-renewal and pluripotency that provides an unlimited resource for research and medical applications. Recently, terminally differentiated murine and human somatic cells were induced to become pluripotent stem (iPS) cells by use of a four-transcription-factor cocktail (Oct-4, Sox-2, Klf4, and Myc; Takahashi and Yamanaka, 2006; Takahashi et al., 2007; Yu et al., 2007; Park et al., 2008). In this unit, the production of retrovirus-expressing reprogramming factors, infection of human fibroblasts, and isolation of human iPS cells will be described. NOTE: The following tissue culture procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All procedures for producing the VSV-G pseudotyped retrovirus should be performed under BL2+ biosafety conditions (according to your Institute’s Safety Department). NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator.

PRODUCTION OF VSV-G PSEUDOTYPED RETROVIRUS This protocol is used for making retroviral vectors pseudotyped with the VSV-G (vesicular stomatitis virus G) envelope protein. The VSV-G pseudotyped retrovirus can be divided into aliquots and stored long-term at −80◦ C and then thawed and used to infect human fibroblasts.

BASIC PROTOCOL 1

Materials 293T cells (ATCC, cat. no. CRL11268) Fugene 6 (Roche Applied Science, cat. no. 1181509001) Current Protocols in Stem Cell Biology 4A.1.1-4A.1.8 Published online January 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc04a01s8 C 2009 John Wiley & Sons, Inc. Copyright 

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DMEM DMEM/F12 (Invitrogen) pMIG-OCT4 (Addgene, clone 17225), pMIG-SOX2 (Addgene, clone 17226), pMIG-KLF4 (clone 17227), and pMIG-MYC (requested from Dr. Cleveland from the Scripps Research Institute) VSV-G (Addgene, clone 8454), and Gag-Pol (Addgene, clone 8455) 293T cell medium (see recipe) 10-cm dishes 0.45-μm filter 38.5-ml polyallomer centrifuge tube (Beckman, cat. no. 326823) Cryovials Additional reagents and equipment for determining the titer of the virus (Sastry et al., 2002; Tiscornia et al., 2006) Transfect 293T cells with plasmids using Fugene 6 1. One the day before transfection, split 293T cells (0.5 × 105 cells/cm2 ) into ten 10-cm dishes for each different virus at a confluency of 40%, aiming to have 70% to 80% confluency the next day. 2. For each 10-cm dish, add 20 μl of Fugene to 300 μl DMEM and incubate the mixture 15 min at room temperature. Add 2.5 μg of pMIG vector, 2.25 μg of Gag-Pol, and 0.25 μg of VSV-G vector and incubate for 15 min. For ten 10-cm dishes, multiply the amounts by 10 to have 3 ml of DMEM with 200 μl Fugene, 25 μg pMIG vector, 22.5 μg Gap-Pol and 2.5 μg VSV-G. Use polystyrene tubes for maximum transfection.

3. While the Fugene/plasmid mixture of step 2 is incubated, aspirate old medium from 293T cells and add 9 ml of new 293T medium. 4. Add 7 ml of 293T cell medium into 3 ml of Fugene/plasmid mixture (for ten 10-cm dishes from step 2) to make a total of 10 ml, mix well by pipetting and add 1 ml of each into each 10-cm dish of 293T cells in 9 ml medium. 5. Place the transfected 293T cells into BL2+ incubator. After 293T cells are transfected with retroviral vectors, they need to be treated as BL2+ hazardous biomaterial.

Concentrate VSV-G pseudotyped retrovirus 6. Three days after transfection (do not change medium during 3-day incubation period), collect and filter the viral supernatant using 0.45-μm filter into a 38.5-ml polyallomer centrifuge tube. 7. Weigh the supernatant tube to make a balance tube for ultracentrifugation. 8. Centrifuge the supernatant 90 min at 70,000 × g, 4◦ C. 9. Remove the supernatant. A white pellet should be visible when a polyallomer centrifuge tube is used.

10. Add 1 ml DMEM/F12 and flick the tubes before storing overnight at 4◦ C to dissolve the pellet. 11. Next day, mix virus by pipetting up and down slowly, aliquot ∼100 to 200 μl of virus into cryovials and store at −80◦ C for long-term storage. Human iPS Cell Derivation/ Reprogramming

12. Determine the titer of the virus following published protocols (Sastry et al., 2002; Tiscornia et al., 2006).

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INFECTION OF FIBROBLASTS AND ISOLATION OF iPS CELLS This protocol is used to infect fibroblasts with the virus, and to isolate iPS cell colonies from them. During and after isolation, human iPS cells show similar colony morphology and require the same culture conditions as human ES cells. Therefore, it is highly recommended that scientists wishing to isolate and maintain human iPS cells first become skilled in handling hES cells, including the mechanical picking and passaging of colonies (UNIT 1C.1), or undergo training directly on iPS cell culture from experienced investigators.

BASIC PROTOCOL 2

Materials Human fibroblasts, acquired from skin biopsy (refer to UNIT 1C.7; split fibroblasts when they reach 70% confluency) Human fibroblast medium (see recipe) Protamine sulfate (see recipe) Retroviral supernatants bearing the appropriate plasmids (Basic Protocol 1) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Mediatech, cat. no. 21-040-CV) MEF (mouse embryonic fibroblasts), CF-1 strain, irradiated (Global Stem, cat. no. GSC-6001G) MEF medium (see recipe) 0.1% (w/v) gelatin (see recipe) 0.05% trypsin/EDTA hESC medium (see recipe) Gelatin-coated 12-well plate preplated with MEFs at a density of 1 × 104 cells/cm2 Collagenase IV (see recipe) Gelatin-coated 6-well plate preplated with MEFs Freezing medium (see recipe) Liquid nitrogen 6-well plate 10-cm dish Dissection microscope 20- and 1000-μl pipets 21-G needle or cell lifter (Corning, cat. no. CT-3008) 15-ml conical tube Cryovials −80◦ C freezer Infect human fibroblasts with retrovirus 1. At a time point 8 to 12 hr prior to infection, plate 1 × 105 human fibroblasts obtained from a skin biopsy in one well of a 6-well plate. 2. Aspirate medium to remove dead cells and add 2 ml of fresh human fibroblast medium. Add protamine sulfate at a final concentration of 5 μg/ml. 3. Into the fibroblast culture, add aliquots of the four retroviral supernatants (Basic Protocol 1) at a multiplicity of infection (MOI) of 5 for each virus. This step and those following must be performed under BL2+ safety conditions.

4. One day after infection, remove the viral supernatant, wash three times, each time with 3 ml PBS, and add 3 ml human fibroblast medium. 5. After 2 more days, replenish dish with 3 ml human fibroblast medium. 6. Four days after infection, plate 1 × 104 /cm2 MEFs in MEF medium on a 10-cm dish coated with 0.1% gelatin. Incubate until the next day.

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A

C

E

B

D

F

Figure 4A.1.1 Identification of iPS colonies among heterogeneous types of colonies arising from infected human fibroblasts. Examples of colonies that should yield stable iPS colonies (A, B) and examples of colonies that do not (C, D). Silencing of retroviral gene expression in iPS colony (E, F). (A) iPS colony with a morphology comparable to hES cells. (B) iPS colony that is surrounded by outgrowth of infected fibroblasts. (C) Non-iPS colony from transformed cells. (D) Non-iPS colony from transformed cells. (E) iPS cell colony in the middle of field (arrow). (F) A reprogrammed iPS colony in E shows no fluorescence due to silencing of the infected retrovirus (arrow). Scale bar = 100 μm.

7. Five days after infection, incubate the infected fibroblasts with 1 ml 0.05% trypsin/EDTA for 3 min at 37◦ C. Stop the trypsinization with 11 ml human fibroblast medium and place all cells into one 10-cm dish preplated with MEFs (prepared in step 6). 8. Seven days after infection remove human fibroblast medium and add 10 ml hESC medium (containing KOSR and 10 ng/ml of bFGF). 9. Replenish the cells with 10 ml hESC medium every day and observe the cells for any sign of colony formation. Typically, sometime around three weeks after infection, various types of colonies will appear, as exemplified in Figure 4A.1.1.

10. Under a dissection microscope and using a 20-μl pipet, pick a colony that shows a morphology similar to hES cells and put the colony into one well of a 12-well plate that has been preplated with MEFs at a density of 1 × 104 cells/cm2 on 0.1% gelatin-coated wells. Fully reprogrammed iPS cells infected with GFP-containing viruses will lack GFP expression, as visualized under a fluorescence microscope (Fig. 4A.1.1F). Absence of GFP expression, a consequence of transcriptional silencing of retroviruses, is specific to embryonic or pluripotent cell types, and can be used as a surrogate marker to identify human iPS cells.

Human iPS Cell Derivation/ Reprogramming

Maintain and store human iPS cells Isolated clones of human iPS cells can be treated and maintained in a similar way as human ES cells. Please refer to the detailed protocol UNIT 1C.1.

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11. When iPS cell colonies grow and start to touch other colonies, wash plate with 1 ml of DMEM/F12, add 0.5 ml collagenase IV, and incubate 10 min at 37◦ C. 12. Using appropriate tools (e.g., 21-G needle, or cell lifter), cut colonies into small pieces, detach pieces from the plate, collect with a 1000-μl pipet (similar to mechanical splitting of established human ES cells; UNIT 1C.1) into a 15-ml conical tube and centrifuge for 4 min at 200 × g, room temperature. Remove the supernatant and add 5 ml of fresh DMEM/F12 followed by centrifuging 4 min at 200 × g, room temperature. 13. Remove the supernatant. 14. For passaging of iPS cells, resuspend pieces of colonies from step 13 in 2 ml fresh hESC medium and transfer into one well of a gelatin-coated 6-well dish precoated with MEFs. Repeat steps 10 to 14 to expand iPS cells. The split ratio used for the cells depends on cell density (ratio is usually 1:3 to 1:6) When you have more than three wells of iPS cells in a 6-well plate, freeze down cells before starting further analysis of iPS cells.

Freeze cells 15. For freezing iPS cells, resuspend colonies from step 13 in 0.5 ml fresh hESC medium, add the same amount of 2× freezing medium dropwise and mix by gently pipetting up and down. Aliquot cells into cryovials and store overnight at –80◦ C. Transfer to liquid nitrogen next day. REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

293T cell medium DMEM containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C Collagenase IV, 10× Dissolve collagenase IV (Invitrogen) at 10 mg/ml in DMEM/F12 (Invitrogen). Filter using 0.22-μm filter. Divide into 0.5- to 1.5-ml aliquots and store up to 1 year at −20◦ C. Before splitting hES or iPS cells, dilute 10× stock solution in DMEM/F12 to make a 1× working stock.

Freezing medium, 2× Make a solution containing 20% dimethyl sulfoxide (DMSO), 60% FBS, and 20% human ES cell (hESC) medium (see recipe). Store up to 1 month at 4◦ C. Gelatin, 0.1% (w/v) Dissolve 0.5 g of gelatin (from porcine skin) in 500 ml distilled water and autoclave. Store indefinitely at room temperature. To gelatinize plates: Prior to addition of MEFs, coat all dishes or wells with enough 0.1% (w/v) gelatin solution to cover the surface. Remove gelatin after 5 min.

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Human embryonic cell (hESC) medium DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (KOSR; Invitrogen) 10 mM non-essential amino acids 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) 50 mM 2-mercaptoethanol 10 ng/ml bFGF (see recipe) Store up to 1 week at 4◦ C Human fibroblast medium MEM-alpha containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C MEF medium DMEM containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C Protamine sulfate, 1000× Dissolve protamine sulfate (Sigma-Aldrich) at 5 mg/ml in phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Mediatech, cat. no. 21-040-CV). Filter using a 0.22-μm filter and store up to 1 year at 4◦ C for future use.

Recombinant human basic fibroblast growth factor (bFGF) Resuspend lyophilized bFGF (PeproTech) to a final concentration of 10 μg/ml in CMF-PBS containing 0.1% (w/v) bovine serum albumin (BSA) and 1 mM DTT. Store at −80◦ C according to manufacturer’s instructions.

COMMENTARY Background Information

Human iPS Cell Derivation/ Reprogramming

Human embryonic stem cells provide a valuable resource for research and regenerative medicine. However, human ES cells isolated to date are not matched to individual patients, and thus allow for generic studies but are limited in their relevance to specific diseases or treatments. Attempts to generate patientspecific stem cells include somatic cell nuclear transfer (SCNT), somatic cell fusion with pluripotent cells, direct cultural adaptation of germ cells, and direct reprogramming of somatic cells with defined factors (Jaenisch and Young, 2008). Each of these approaches has specific advantages and limitations. Since the report by Yamanaka’s group that mouse embryonic and adult tail-tip fibroblasts can be reprogrammed to become ES cell–like pluripo-

tent cells by expression of four transcription factors (Oct4, Sox2, Klf4, c-Myc), a similar strategy has been used to isolate human iPS cells. Our laboratory and Yamanaka’s group successfully isolated human iPS cells from embryonic and adult human fibroblasts using the same four transcription factors (Takahashi et al., 2007; Park et al., 2008). A different combination of factors (Nanog and Lin28 in place of Klf4 and Myc) also produced human iPS cells (Yu et al., 2007). Yamanaka’s reprogramming strategy also works even without Myc, although at lower efficiency, to produce iPS cells from both murine and human fibroblasts (Nakagawa et al., 2008). The isolation of murine iPS cells has been facilitated by using fibroblasts that carry endogenous selectable markers. Fibroblasts from

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mice that express neomycin- or puromycinresistance genes under the control of the promoters for Fbx15, Oct4, or Nanog loci have been infected with four reprogramming factors, selected with neomycin or puromycin, and successfully reprogrammed to become iPS cells (Takahashi and Yamanaka, 2006; Maherali et al., 2007; Okita et al., 2007; Wernig et al., 2007). For most human fibroblasts, such endogenous selection systems do not exist, and thus selection has been based mostly upon colony morphology alone. By selecting colonies that show similar morphology to human ES cells, human iPS cells can be readily selected from a morphologically diverse background of colonies that arise when human fibroblasts are infected with several retroviruses. Although reprogramming human fibroblasts into pluripotent cells will provide an alternative to human ES cells for certain research and clinical applications, current methods of generating iPS cells employ retroviral vectors that integrate into the fibroblast genome, and thus the resulting cells are potentially tumorigenic. Developing virus-free methods is desirable. Excision of ectopic Myc by Cremediated recombination has been shown to reduce the tumor formation potential of iPS cells (Hanna et al., 2007; Shi et al., 2008). The G9a inhibitor BIX01294 can replace Oct4, and only two factors are sufficient to make iPS cells from murine neuronal stem cells (Shi et al., 2008), suggesting that the right combination of excisable viruses and chemicals may provide a method to make iPS cells that lack persistent viral integration.

Critical Parameters and Troubleshooting Before starting to reprogram human fibroblasts, there are three important factors that determine success: the quality of the virus, MEFs, and target fibroblasts. When a new VSV-G pseudotyped virus is made and stored at –80◦ C, be sure to determine the titer of the virus, because it is essential to have a highquality, high-titer virus. If the titer is <0.5 × 106 IU/ml, make a new virus. The proper storage of the virus is important to maintain quality and titer, and we recommend freezing a large number of small aliquots of the virus, and limiting thawing and refreezing. If the virus has been stored for a long period of time, determine the titer again before starting another round of reprogramming. The quality of MEFs is also an important factor. Con-

firm the quality of the lot of MEFs by culturing hES cells on them before plating infected human fibroblasts. When low-quality MEFs are used to select iPS cells, the initial colonies of reprogrammed fibroblasts differentiate and do not form human ES cell-like colonies. The quality of human fibroblasts is another important factor to determine the efficiency of reprogramming. Although the mechanism is not yet clear, fibroblasts from younger donors appear to yield more iPS colonies than those from older donors. Beginners are recommended to start reprogramming with fetal or foreskin fibroblasts to experience the procedures of isolating iPS cells before attempting reprogramming of fibroblasts from older individuals.

Anticipated Results The efficiency of reprogramming human fibroblasts depends on parameters described in the troubleshooting section. By using a highquality virus and MEFs, this protocol generates ∼5 to 50 colonies from 1 × 105 cells, for a frequency of ∼0.05%. When iPS cell lines are formed, they can be passaged over a year without karyotypic abnormality.

Time Considerations Two weeks are required to make a retrovirus carrying the reprogramming factors, including thawing of the 293T cells, transfecting of vectors, and determining of titer for the virus. While the retrovirus is being made, test the quality of the MEFs by culturing human ES cells on them. Three to four weeks are required to identify and isolate iPS cells, beginning from the day human fibroblasts are infected. Single iPS colonies need to be expanded for two to three weeks to generate stable cell lines that have an adequate number of cells for subsequent characterization.

Acknowledgements Georg Q. Daley was supported by grants from the National Institutes of Health, the NIH Director’s Pioneer Award of the NIH Roadmap for Medical Research, and private funds contributed to the Harvard Stem Cell Institute and the Children’s Hospital Stem Cell Program. George Q. Daley is a recipient of Clinical Scientist Awards in Translational Research from the Burroughs Wellcome Fund and the Leukemia and Lymphoma Society, and is an Investigator of the Howard Hughes Medical Institute.

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Literature Cited Hanna, J., Wernig, M., Markoulaki, S., Sun, C.W., Meissner, A., Cassady, J.P., Beard, C., Brambrink, T., Wu, L.C., Townes, T.M., and Jaenisch, R. 2007. Treatment of sickle cell anemia mouse model with iPS cells generated from autologous skin. Science 318:19201923. Jaenisch, R. and Young, R. 2008. Stem cells, the molecular circuitry of pluripotency and nuclear reprogramming. Cell 132:567-582. Maherali, N., Sridharan, R., Xie, W., Utikal, J., Eminli, S., Arnold, K., Stadtfeld, M., Yachechko, R., Tcheiu, J., Jaenisch, R., Plath, K., and Hochedlinger, K. 2007. Directly reprogrammed fibroblasts show global epigenetic remodeling and widespread tissue contribution. Cell Stem Cell 1:55-70. Nakagawa, M., Koyanagi, M., Tanabe, K., Takahashi, K., Ichisaka, T., Aoi, T., Okita, K., Mochiduki, Y., Takizawa, N., and Yamanaka, S. 2008. Generation of induced pluripotent stem cells without Myc from mouse and human fibroblasts. Nat. Biotechnol. 26:101-106. Okita, K., Ichisaka, T., and Yamanaka, S. 2007. Generation of germline-competent induced pluripotent stem cells. Nature 448:313-317. Park, I.H., Zhao, R., West, J.A., Yabuuchi, A., Huo, H., Ince, T.A., Lerou, P.H., Lensch, M.W., and Daley, G.Q. 2008. Reprogramming of human somatic cells to pluripotency with defined factors. Nature 451:141-146.

Sastry, L., Johnson, T., Hobson, M.J., Smucker, B., and Cornetta, K. 2002. Titering lentiviral vectors: Comparison of DNA, RNA and marker expression methods. Gene Ther. 9:1155-1162. Shi, Y., Do, J.T., Desponts, C., Hahm, H.S., Scholer, H.R., and Ding, S. 2008. A combined chemical and genetic approach for the generation of induced pluripotent stem cells. Cell Stem Cell 2:525-528. Takahashi, K., Tanabe, K., Ohnuki, M., Narita, M., Ichisaka, T., Tomoda, K., and Yamanaka, S. 2007. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861-872. Takahashi, K. and Yamanaka, S. 2006. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126:663-676. Tiscornia, G., Singer, O., and Verma, I.M. 2006. Production and purification of lentiviral vectors. Nat. Protoc. 1:241-245. Wernig, M., Meissner, A., Foreman, R., Brambrink, T., Ku, M., Hochedlinger, K., Bernstein, B.E., and Jaenisch, R. 2007. In vitro reprogramming of fibroblasts into a pluripotent ES-cell-like state. Nature 448:318-324. Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S., Nie, J., Jonsdottir, G.A., Ruotti, V., Stewart, R., Slukvin, I.L., and Thomson, J.A. 2007. Induced pluripotent stem cell lines derived from human somatic cells. Science 318:1917-1920.

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Generation and Characterization of Human Induced Pluripotent Stem Cells

UNIT 4A.2

Mari Ohnuki,1, 2 Kazutoshi Takahashi,1 and Shinya Yamanaka1, 2, 3, 4 1

Institute for Integrated Cell-Material Sciences, Kyoto University, Kyoto, Japan Institute for Frontier Medical Sciences, Kyoto University, Kyoto, Japan 3 Japan Science and Technology Agency, Kawaguchi, Japan 4 Gladstone Institute of Cardiovascular Disease, San Francisco, California 2

ABSTRACT This unit describes how to generate human induced pluripotent stem (iPS) cells and evaluate the qualities of the generated iPS cells. The methods for establishment and maintenance of human iPS cells are similar to those for mouse iPS cells but not identical. In addition, these protocols include excellent procedures for passaging and cryopreservation of human iPS cells established by ES cell researchers, which result in an easy way to culture human iPS cells. Moreover, we include methods for characterizing iPS cells for further research. RT-PCR and immunocytochemistry for detection of pluripotent cell markers, embryoid body differentiation, and teratoma differentiation are used to determine pluripotency in vitro and in vivo, respectively. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 9:4A.2.1-4A.2.25.  Keywords: reprogramming r pluripotency r iPS cells

INTRODUCTION The authors of this unit have reported that the forced expression of four transcription factors, Oct3/4, Sox2, Klf4, and c-myc, could reprogram fibroblasts to pluripotent stem cells (Takahashi and Yamanaka, 2006; Yamanaka, 2007). These reprogrammed cells are referred to as induced pluripotent stem (iPS) cells. By using Nanog or Oct3/4 as selection markers, we and others have successfully induced germ line competency with these four factors (Maherali et al., 2007; Okita et al., 2007; Wernig et al., 2007). Recently, it was demonstrated that drug selection with pluripotent stem cell markers is not required for establishment of iPS cells (Blelloch et al., 2007; Meissner et al., 2007; Nakagawa et al., 2008). Reprogrammed cells formed round-shaped colonies and could be morphologically distinguished by microscopic observation. The result suggests that iPS cells can be established utilizing somatic cells from genetically unmodified animals, and provides hope of medical applications in human cells with genes that are hard to modify. The same set of four factors allowed reprogramming human adult fibroblasts to the pluripotent state (Takahashi et al., 2007a; Yu et al., 2007; Lowry et al., 2008; Masaki et al., 2008; Park et al., 2008a). Generation of human iPS cells requires only basic techniques in molecular and cell biology and does not require any special equipment. In this unit, we introduce not only how to generate iPS cells but also how to evaluate the characteristics of iPS cells. The methods for establishment and maintenance of iPS cells are similar to those for mouse iPS cells, but not identical (Takahashi et al., 2007b; Basic Protocol 1). Support Protocols describe preparation of SNL feeder cells (Support Protocol 1) and preparation of PLAT-E packaging cells (Support Protocol 2). In addition, our protocols include excellent procedures for passaging (Basic Protocol 2) and cryopreservation (Basic Protocol 3) of human iPS cells established by ES cell researchers Current Protocols in Stem Cell Biology 4A.2.1-4A.2.25 Published online June 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc04a02s9 C 2009 John Wiley & Sons, Inc. Copyright 

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(Fujioka et al., 2004; Watanabe et al., 2007), which results in an easy way to culture human iPS cells. Moreover, we demonstrate the methods to characterize iPS cell clones for further in-depth research. RT-PCR for detection of pluripotent cell markers (Support Protocol 3), immunocytochemistry for pluripotent cell markers (Support Protocol 4), and in vitro and in vivo differentiations by embryoid body (Support Protocol 5) and teratoma formation (Support Protocol 6) are extensively described. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. NOTE: All incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. BASIC PROTOCOL 1

GENERATION OF iPS CELLS The first step in generation of iPS cells is to transduce mouse ecotropic retrovirus receptor genes into human skin fibroblasts. These genes are necessary to enhance transduction efficiency of transgenes and increase safety for the experimenters. The next step is to introduce the factors to be tested for their ability to induce iPS cells. See Figure 4A.2.1 for an outline of the procedure. For gene transduction, we utilized the combination of pMXs retroviral vector and PLAT-E packaging cells, which can produce a much higher titer of retrovirus, serving as a sufficient vector to generate mouse iPS cells. Despite the fact that ecotropic retrovirus that affects exclusively rodent cells is produced, we decided to apply this combined system to human cells for the safety of research personnel. In order to enable ecotropic retroviruses to transduce human cells, we introduced mouse solute carrier family 7 (cationic amino acid transporter, y+ system) member 1 (Slc7a1) gene encoding the ecotropic retrovirus receptor into human cells. We use SNL cells as feeder cells for maintenance of both mouse and human iPS cells. These cell lines are derived from mouse embryos and express the neomycin resistance gene and the leukemia inhibitory factor (LIF) gene. These cell lines provide two significant merits: they show no remarkable difference between tubes and they have a more extended period of proliferation when compared to primary mouse embryonic fibroblasts (MEFs). We have always used iPS cells before passage 20, but the highest limit on passage number is not known. CAUTION: All processes involving lentivirus should be performed in a safety cabinet while wearing gloves. All waste must be treated with first with ethanol, then with bleach (hypochlorous acid), and finally autoclaved.

Materials

Generation and Characterization of Human iPS Cells

293FT cells for producing lentivirus (Invitrogen; see manufacturer-provided protocol for culture) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) 0.25% trypsin/1 mM EDTA solution (Invitrogen, cat. no. 25200-056) 293FT medium (see recipe) OPTI-MEM I medium (Invitrogen, cat. no. 31985-062) ViraPower packaging mix (from ViraPower expression system kit; Invitrogen, cat. no. K4990-00) pLenti6/UbC containing mouse Slc7a1 gene (Addgene; http://www.addgene.org/ Shinya Yamanaka) Lipofectamine 2000 (Invitrogen, cat. no. 11668-019)

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Figure 4A.2.1

timing d0

condition

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4A.2.3

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non-iPS (non-ES-like) colony iPS (ES-like) colony

fibroblast transduced with retroviruses

d60~

human fibroblast expressing mouse Slc7a1

d45~

SNL cells (mitomycin c–treated)

d40

human fibroblast

d30

Schematic diagram of iPS cell generation. A strategy and approximate time table for human iPS cell generation.

d15

ES-like cells arrive late

d85~

establishment of iPS cell line

picking passaging the iPS colonies and cryopreserving

ES medium + bFGF

appearance of non-ES-like colonies

replating fibroblasts onto SNL feeder

preparation of SNL feeder

FP medium (10% FBS)

preparation of fibroblasts

lentivirus production and infection

retrovirus production and infection

10% FBS medium (see recipe) Human fibroblast cells (available from the following sources: Cell Applications Inc. (http://www.cellapplications.com/) Lonza (http://www.lonza.com/group/en.html) American Type Culture Collection (ATCC, http://www.atcc.org/) European Collection of Cell Cultures (ECACC; http://www.ecacc.org.uk/) Riken Bioresource Center (http://www.brc.riken.jp/) Japanese Collection of Research Bioresources (http://cellbank.nibio.go.jp/) 0.05% trypsin/0.53 mM EDTA solution (Invitrogen, cat. no. 25300-054) Hexadimethrine bromide (polybrene; Nacalai Tesque, cat. no. 17736-44) PLAT-E packaging cells (Support Protocol 2) pMXs retroviral vectors encoding OCT3/4, SOX2, KLF4, and/or c-myc (Addgene; http://www.addgene.org/Shinya Yamanaka): pMXs-hOCT3/4 pMXs-hSOX2 pMXs-hKLF4 pMXs-hc-MYC pMXs retroviral vector encoding the green fluorescence protein (GFP) to monitor transfection efficiency and serve as a negative control for iPS cell induction (Cell Biolabs, Inc.) Fugene 6 transfection reagent (Roche, cat. no. 1 814 443) Mitomycin C–treated SNL feeder cell plates, 100-mm and 24-well (Support Protocol 1) hES cell medium (see recipe) Recombinant basic fibroblast growth factor, human (bFGF; Wako, cat. no. 064-04541) 100-mm tissue culture dish (Falcon, cat. no. 353003) 0.45-μm pore size cellulose acetate filter (Whatman, cat. no. FP30/0.45 CA-S) 96-well tissue culture plate (Falcon, cat. no. 351172) 24-well tissue culture plate (Falcon, cat. no. 353047) Additional reagents and equipment for counting cells (UNIT 1C.3) and preparation of PLAT-E packaging cells (Support Protocol 2) Passage 293FT cells 1. Aspirate the medium from an 80% to 90% confluent culture of 293FT cells growing in a 100-mm tissue culture dish and wash the cells once with 10 ml of CMFDPBS. Add 1 ml of 0.25% trypsin/1 mM EDTA and incubate for 2 min at room temperature. 2. Add 9 ml of 293FT medium and break the cells into single-cell suspension by pipetting up and down about 10 times. 3. Determine cell number using a hemacytometer (UNIT 1C.3), plate 4 × 106 cells on 100-mm dish and incubate overnight at 37◦ C, in a humidified 5% CO2 incubator.

Prepare virus 4. Dispense 1.5 ml of OPTI-MEM I medium into a 1.7-ml tube. Add 9 μg of ViraPower packaging mix (including pLP1, pLP2, and pLP/VSVG) and 3 μg of pLenti6/UbC encoding mouse Slc7a1 gene, and mix them gently. 5. In another 1.7-ml tube, dispense 1.5 ml of OPTI-MEM I medium and add 36 μl of Lipofectamine 2000, and mix gently. Incubate for 5 min at room temperature. Generation and Characterization of Human iPS Cells

6. Mix Lipofectamine 2000 diluted solution and previous DNA mixture gently and incubate for 20 min at room temperature to form a DNA/Lipofectamine complex.

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Produce virus 7. Replace the medium on 293FT cells (see step 3) with fresh 10% FBS medium. 8. Add 3 ml of DNA/Lipofectamine complex (from step 6) to the dish of 293FT cells and rock it back and forth gently. Incubate the dish overnight in a 37◦ C, 5% CO2 incubator. 9. At a time point 24 hr after transfection, replace the medium with 10 ml of fresh 10% FBS medium. Incubate the dish for another 24 hr.

Begin culturing human fibroblasts 10. In preparation for infection, plate 5 × 105 human fibroblasts in 100-mm dish with 10 ml of 10% FBS medium. Incubate. Collect virus 11. On the next day, collect the virus-containing supernatant from the 100-mm dish of transfected 293FT cells with a 10-ml disposable syringe. Filter the supernatant with a 0.45-μm pore size cellulose acetate filter. 12. Use the virus-containing medium immediately (see step 17) or store at −80◦ C.

Prepare fibroblasts Fibroblasts should be replated as described below on the day before infection for the sake of better infection efficiency. 13. Remove the 100-mm dish of human fibroblasts (step 10) from the incubator. 14. Aspirate the medium and wash the cells once with 10 ml CMF-DPBS, add 1 ml of 0.05% trypsin/0.53 mM EDTA, and incubate for 10 min at 37◦ C. 15. Add 9 ml of 10% FBS medium and break the cells into a single-cell suspension by pipetting. 16. Determine cell number (UNIT 1C.3), plate 8 × 105 cells on a 100-mm dish, and incubate overnight.

Infect the cells 17. Replace the medium on the fibroblasts with the virus cocktail (entire supernatant from step 11) supplemented with 4 μg/ml polybrene (hexadimethrine bromide). Incubate the dish 5 hr to overnight. 18. After infection, wash the cells with 10 ml of CMF-DPBS (optional), and exchange the medium with 10 ml of fresh 10% FBS medium at room temperature. Return the fibroblast cultures to the incubator. Sometimes overnight incubation with lentivirus is toxic to fibroblasts. In that case, dilute virus cocktail by ∼50% with medium or shorten the incubation time to 5 hr.

19. To check the expression of infected genes, use a GFP-encoding vector as a control. Infection can also be confirmed by culturing in blasticidin S–supplemented medium (10 μg/ml) because pLenti6/UbC/mSlc7a1 includes the blasticidin S–resistance gene. Expression is determined by microscopic examination or flow cytometry.

Prepare retroviruses 20. Prepare a single-cell suspension of PLAT-E cells (Support Protocol 2). The protocol is based on 100-mm dish cultures of cells. If you use different sizes of dishes or plates, adjust the cell numbers and volumes according to Table 4A.2.1.

21. Transfer 3.6 × 106 PLAT-E cells to a new 100-mm dish in 10% FBS medium without puromycin or blasticidin S. Prepare one dish per plasmid (the plasmids will be pMXs

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Table 4A.2.1 Amounts of Reagents for Generating Plasmids in Different Size Culture Dishes

Reagents

100-mm dish

60-mm dish

6-well plate

PLAT-E cells

3.6 × 106

1.2 × 106

6 × 105

OPTI-MEM I

300 μl

100 μl

50 μl

Fugene 6

27 μl

9 μl

4.5 μl

Plasmid

9 μg

3 μg

1.5 μg

encoding OCT3/4, SOX2, KLF4, and/or c-myc, as well as GFP as a control). Incubate the dishes overnight. One plasmid DNA involves one dish of PLAT-E cells and, consequently, introducing genes for four factors and the GFP control requires five dishes of PLAT-E cells.

22. The day after passage of PLAT-E cells, prepare one 1.5-ml microcentrifuge tube per plasmid DNA. 23. Dispense 0.3 ml of OPTI-MEM I into each tube. 24. Add 27 μl of Fugene 6 transfection reagent into each tube of OPTI-MEM I and mix gently with finger tapping. Incubate tubes for 5 min at room temperature. 25. Add 9 μg of the appropriate plasmid DNA to each tube, one plasmid per tube. Mix by tapping and incubate tubes for 15 min at room temperature. 26. Add each DNA/Fugene 6 mixture to one of five separate cultures of PLAT-E cells (see step 21). Incubate the dishes overnight. Monitor the efficiency of transfection with GFP-coding pMXs vector. Our laboratory confirms transduction efficiency of more than 60%. High efficiency is essential for iPS induction.

27. On the next day, replace the medium containing DNA and Fugene with fresh 10% FBS medium and return the dishes to the incubator. 28. The day after transfection of the PLAT-E cells, prepare a suspension of mouse Slc7a1-expressing human fibroblasts in 10% FBS medium, as described in steps 13 to 16. Count cells and plate 8 × 105 cells per 100-mm dish (see step 16), and incubate the dish overnight. 29. On the next day, collect the virus-containing medium from each of the dishes of transfected PLAT-E cells with a 10-ml disposable syringe and filter it with a 0.45-μm pore size cellulose acetate filter. 30. Mix equal amounts of each of the three or four virus-containing media (OCT3/4, SOX2, KLF4, with or without c-myc). The virus cocktail should be applied immediately. Do not freeze, or infection efficiency will be lower. The virus with the GFP control vector will be applied to a separate plate of human fibroblasts.

Generation and Characterization of Human iPS Cells

31. Replace the medium on fibroblasts expressing Slc7a1 with the virus cocktail (step 30) supplemented with 4 μg/ml polybrene. Incubate the dish for 4 hr to overnight. 32. At a time point 24 hr after infection, change the medium to 10 ml of fresh 10% FBS medium. Change the medium every second day until reseeding.

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Figure 4A.2.2 Images related to iPS cell induction. (A, B) Images of non-iPS and ES-like (iPS) cells, respectively. (C) Image of established iPS cells. Images of immunocytochemistry for undifferentiated pluripotent cell markers: (D) SSEA3; (E) TRA-1-60; (F) TRA-1-81; (G) Nanog; and (H) SSEA1 (negative). Blue indicates nuclei stained with Hoechst 33342. Images of immunocytochemistry for differentiated cell products: (I) α-fetoprotein; (J) α-smooth muscle actin; and (K) βIII-tubulin. Blue indicates nuclei stained with Hoechst 33342. (L) Image of SCID mouse that had iPS cells injected into the testes 3 months earlier. (M) Image of a dissected teratoma. (N) Image of hematoxylin and eosin–stained teratoma section. Bars = 100 μm. Panels D to H illustrate immunohistochemistry for pluripotent cell markers, while panel I illustrates a marker for endoderm, panel J a marker for mesoderm, and panel K a marker for ectoderm.

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Reseed fibroblasts on SNL feeder cells 33. At a time point 6 days after infection, aspirate the medium and wash the cells once with 8 ml of CMF-DPBS. 34. Add 1 ml of 0.05% trypsin/0.53 mM EDTA and incubate the dish 10 min at 37◦ C for 10 min. 35. Add 9 ml of 10% FBS medium to the dish and break up the mass of cells by pipetting. 36. Determine the cell number (UNIT 1C.3) and transfer 5 × 104 or 5 × 105 cells to a 100mm dish with mitomycin C–treated SNL feeder cells (Support Protocol 1). Incubate the dish overnight. 37. The next day and every second day, change the medium to 10 ml of hES cell medium supplemented with 4 ng/ml bFGF.

Pick colonies 38. At a time point 2 to 3 weeks after retroviral infection, examine the dishes for colonies (see Fig. 4A.2.2). It takes about 30 days for iPS cell colonies to grow large enough to be picked up. Observe the dishes carefully because the timing of colony emergence differs in each experiment even if the same fibroblast clones were induced. Fuzzy-edged colonies appear ∼2 weeks after infection; these are not the iPS colonies. Wait another week before ES cell-like, clear-edged colonies begin to be seen; these are the iPS cell colonies that should be picked.

39. Distribute 20 μl of hES cell medium to each well of a 96-well plate. Wash the dish of iPS cell colonies once with 10 ml of CMF-DPBS, and then add another 5 ml of CMF-DPBS. 40. With the 5 ml of CMF-DPBS still in the dish, cut out an iPS colony and separate it from feeder cells under the stereo microscope with a 2- or 10-μl pipet tip and pipettor. Transfer the colony to an individual well of 96-well plate. 41. After picking the colonies add 180 μl of hES medium and break the colonies into small masses of cells but not single cells by pipetting. 42. Transfer the suspension to a well of a 24-well plate with SNL feeder cells. Incubate the plate until the cells grow to 80% to 90% confluency. Continue passaging as in Basic Protocol 2. SUPPORT PROTOCOL 1

PREPARATION OF SNL FEEDER CELLS SNL cells are mitomycin C inactivated and used as feeder cells for plating iPS cells. We have always used feeder cells before passage 20, but the highest limit on passage number is not known.

Materials

Generation and Characterization of Human iPS Cells

Frozen vial of SNL feeder cells (McMahon and Bradley, 1990): available from Dr. Allan Bradley of the Sanger Institute (http://www.sanger.ac.uk/Teams/ faculty/bradley/) SNL medium (see recipe) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) 0.25% trypsin/1 mM EDTA solution (Invitrogen, cat. no. 25200-056) 0.4 mg/ml mitomycin C (see recipe)

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Centrifuge Gelatin-coated 100-mm tissue culture dish (see recipe) Other gelatin-coated culture vessels (see recipe) as needed Additional reagents and equipment for counting cells (UNIT 1C.3) 1. Place a frozen vial of SNL cells in a 37◦ C water bath until almost thawed. Resuspend the cells in 9 ml of SNL medium. 2. Centrifuge 5 min at 160 × g, room temperature, and discard the supernatant. 3. Resuspend the cells in 10 ml of fresh SNL medium, and transfer to a gelatin-coated 100-mm dish (∼1 × 106 cells). Incubate the cells in a humidified 37◦ C, 5% CO2 incubator until the cells are 80% to 90% confluent. Do not make cells overconfluent, or their performance as feeder cells may deteriorate.

4. Aspirate off the medium and wash the cells once with 8 ml of CMF-DPBS. Add 0.5 ml of 0.25% trypsin/1 mM EDTA and incubate for 1 min at room temperature. 5. Add 4.5 ml of SNL medium and break the cells into a single-cell suspension by pipetting up and down several times. 6. Split the cell suspension 1:16, plate on a gelatin-coated 100-mm dishes, and incubate (3 to 4 days) until the cells are 80% to 90% confluent. 7. When the cells reach 80% to 90% confluency, drop 0.3 ml of 0.4 mg/ml mitomycin C solution on the culture of SNL cells and mix by gently shaking back and forth. Incubate 2.25 hr in humidified 37◦ C, 5% CO2 incubator. 8. After incubation, aspirate the mitomycin C–containing medium and wash the cells with 5 ml of CMF-DPBS twice. 9. Add 0.5 ml of 0.25% trypsin/1 mM EDTA and incubate for 1 min at room temperature. Add 4.5 ml of SNL medium and break the cells into a single-cell suspension by pipetting. 10. Count cells (UNIT 1C.3), and plate 1.5 × 106 cells (in 10 ml SNL medium) per gelatincoated 100-mm dish, 2.5 × 105 cells (in 2 ml SNL medium) per well of 6-well plate, or 6.1 × 104 cells (in 0.5 ml SNL medium) per well of a 24-well plate. Cells should be nicely spread with few gaps in between.

11. Incubate the dish overnight. The cells should become ready for use by the next day. SNL feeder cell–plated dishes should be used within 3 days.

PREPARATION OF PLAT-E PACKAGING CELLS PLAT-E packaging cells are used to prepare the viral stocks bearing the plasmids for induction of iPS cells. The PLAT-E packaging cell line is designed for producing ecotropic retrovirus; the cells are derived from HEK293 cells and contain env-IRES-puroR and gag-pol-IRES-bsR cassettes driven by the human elongation factor 1α promoter.

SUPPORT PROTOCOL 2

Materials Frozen vial of PLAT-E packaging cells (Morita et al., 2000): available from Dr. Toshio Kitamura at the University of Tokyo ([email protected]) or Cell Biolabs, Inc. (http://www.cellbiolabs.com/) 10% FBS medium (see recipe) Puromycin stock solution (see recipe)

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Blasticidin S stock solution (see recipe) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) 0.05% (w/v) trypsin/0.53 mM EDTA solution (Invitrogen, cat. no. 25300-054) 100-mm tissue culture dishes Centrifuge 15-ml conical centrifuge tubes 1. Place a frozen vial of PLAT-E packaging cells in a 37◦ C water bath until almost thawed. Resuspend the cells in 9 ml of 10% FBS medium. 2. Centrifuge 5 min at 180 × g, room temperature, and discard the supernatant. 3. Resuspend the cells in 10 ml of fresh 10% FBS medium, and transfer them to a 100-mm dish. Incubate the cells in a humidified 37◦ C, 5% CO2 incubator. 4. On the next day, change the medium to fresh 10% FBS medium supplemented with 1 μg/ml puromycin and 10 μg/ml blasticidin S. Incubate until the cells are 80% to 90% confluent. 5. Aspirate the medium and wash the cells once with 10 ml of CMF-DPBS. Add 4 ml of 0.05% trypsin/0.53 mM EDTA and incubate for 1 min at room temperature. 6. Tap the dish and add 10 ml of 10% FBS medium, then transfer cell suspension to a 15-ml conical tube. 7. Centrifuge 5 min at 180 × g, room temperature, and discard the supernatant. 8. Resuspend in 10 ml of 10% FBS medium and break the mass of cells into single-cell suspension by pipetting. 9. Split the cell suspension 1:4 to 1:6, plate on 100-mm dishes, and incubate (2 to 3 days) until the cells are 80% to 90% confluent. BASIC PROTOCOL 2

PASSAGE OF iPS CELLS The following protocol is based on the cells of 24-well plate cultures. If you use different sizes of dishes or plates, adjust the volume according to Table 4A.2.2.

Materials Human iPS cells at 80% to 90% confluency in a 24-well plate (Basic Protocol 1) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium (see recipe) 6-well plate seeded with mitomycin C–treated SNL cells (Support Protocol 1) Sterile disposable cell scraper 15-ml conical centrifuge tube 1. Wash 24-well plate of 80% to 90% confluent iPS cells once with 0.5 ml per well of CMF-DPBS. 2. Aspirate CMF-DPBS completely, add 0.1 ml of CTK solution to the dish, and incubate for ∼5 min at 37◦ C. Generation and Characterization of Human iPS Cells

3. When ∼90% of feeder cells detach, wash out CTK solution and feeder cells with 0.5 ml of CMF-DPBS, twice. Usually, feeder cells detach first from the dish, whereas iPS colonies remain attached.

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Table 4A.2.2 Reagent Volumes for Passaging iPS Cells Grown in Different Culture Dishes and Plates

Reagent

100-mm

60-mm

6-well

24-well

PBS

10 ml

4 ml

2 ml

0.5 ml

CTK solution

1 ml

0.5 ml

0.3 ml

0.1 ml

hES medium (ml)

10 ml

4 ml

2 ml

0.5 ml

4. Remove CMF-DPBS completely, and add 0.5 ml of hES medium to the dish. 5. Scrape out the iPS colonies by using sterile disposable cell scraper, and break the colonies into small clumps by pipetting up and down. Do not break the colonies up completely into single cells, because too much dissociation might trigger cell death.

6. Transfer the cell suspension to a 15-ml conical tube. 7. Dilute the cell suspension at 1:3 to 1:4 with hES medium, and transfer 2 ml of the suspension to a new well of a 6-well plate seeded with mitomycin C–treated SNL feeders. 8. Incubate in humidified 37◦ C, 5% CO2 incubator. Change the medium with fresh hES cell medium every day. Cells are passaged approximately every 5 days.

STORAGE OF ESTABLISHED iPS CELLS Established cultures of iPS cells should be frozen at early passages to maintain the stock. When the cells reach confluency (i.e., when the colonies approach each other) in the 100-mm dish, it is time to make cryostocks. This method uses a specific inhibitor for p160-Rho-associated coiled-coil kinase (ROCK), Y-27632, to increase the survivability of the frozen cells.

BASIC PROTOCOL 3

Materials 10 mM Y-27632 (Wako, cat. no. 253-00513) Confluent iPS cells (see Basic Protocols 1 and 2) in 100-mm dishes Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium (see recipe) DAP123 solution (see recipe) Liquid nitrogen 60-mm dish seeded with mitomycin C–treated SNL feeder cells (Support Protocol 1) Sterile disposable cell scraper 2-ml cryovials Liquid nitrogen tank Prepare cells for freezing 1. Add 10 μl of 10 mM Y-27632 to the medium of a confluent iPS cell culture in a 100-mm dish, and incubate the dish at 37◦ C for at least 1 hr. 2. Aspirate the medium, and wash the cells with 10 ml of CMF-DPBS.

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3. Discard CMF-PBS, add 1 ml of CTK solution, and incubate at 37◦ C for 2 to 5 min. Incubation time may depend on cell density. Check the cells by eye once per minute. After treatment with Y-27632, the cells may become less detachable. In such cases, you can treat the cells with the CTK solution for a longer period of time (∼10 min).

4. Wash twice, each time with 10 ml of CMF-DPBS to remove feeder cells and CTK solution. 5. Discard CMF-DPBS, add 12 ml of hES medium, detach the colonies from the dish using a cell scraper, and transfer 4 ml of cell suspension to each of three 15-ml conical tubes. 6. Centrifuge 5 min at 160 × g, room temperature. 7. Remove the supernatant.

Freeze the cells 8. Resuspend the pellet in 0.2 ml of DAP213 solution by pipetting a few times with pipettor. Do not break up the colonies.

9. Transfer 0.2 ml of the cell suspension to 2-ml cryovials. 10. Put the vials quickly into liquid nitrogen. After adding DAP213 to the cells, the suspension must be frozen within 15 sec for viability of the cells.

11. Store the cells in the liquid nitrogen tank.

Thaw frozen stock 12. Prepare 10 ml of prewarmed (37◦ C) hES medium in a 15-ml conical tube. 13. Remove frozen vial of iPS cells from liquid nitrogen tank. 14. Add 0.8 ml of prewarmed hES medium to the vial and thaw the cells quickly by pipetting up and down with a 1000-μl pipet tip and pipettor. 15. Transfer the cell suspension to the tube prepared in step 12. 16. Centrifuge 5 min at 160 × g, room temperature. 17. Discard the supernatant, and add 4 ml of hES medium. 18. Transfer the cell suspension to a 60-mm dish seeded with mitomycin C–treated SNL feeder cells and incubate in a humidified 37◦ C, 5% CO2 incubator. For the viability of the iPS cells, steps 14 to 16 should be finished as quickly as possible. Do not break up the cell clumps into single cells. SUPPORT PROTOCOL 3

RT-PCR FOR DETECTION OF PLURIPOTENT CELL MARKERS RT-PCR for marker genes of pluripotent stem cells is one of the easiest assays to evaluate the quality of iPS cells. The expression of not only endogenous genes but also transgenes from retroviruses can be examined by RT-PCR.

Materials Generation and Characterization of Human iPS Cells

Human iPS cells cultured in 6-well plate (Basic Protocols 1 and 2), 80% to 90% confluent Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) Trizol reagent (Invitrogen, cat. no. 15596-026)

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Chloroform (Nacalai Tesque) Isopropanol (Nacalai Tesque) 70% ethanol in nuclease-free water Nuclease-free (e.g., Milli-Q) water Turbo DNA-free Kit (Ambion, cat. no. AM1907) containing: 10× DNase I buffer Recombinant DNase I DNase Inactivation Reagent ReverTra Ace -α- kit (Toyobo, cat no. FSK-101; http://www.toyobo.co.jp/) containing: 5× reverse transcription buffer (containing 25 mM Mg2+ ) 10 mM dNTPs Recombinant ribonuclease inhibitor (10 U/μl) Reverse transcriptase Oligo dT20 primer (10 pmol/μl) ExTaq kit (Takara, cat. no. RR001A; http://www.takara-bio.us) containing: ExTaq DNA polymerase (5 U/μl) 10× ExTaq buffer 2.5 mM dNTPs PCR primers for human ES cell markers (Figure 4A.2.3) 15-ml conical tubes (Falcon) Centrifuge Nanodrop spectrometer (Thermo Scientific) Filtered pipet tips 10-μl, 200-μl, and 1000-μl (RNase-free, Watson) 1.5-ml microcentrifuge tubes, RNase-free 0.2-ml PCR reaction tubes (Greiner) Thermal cycler (Applied Biosystems) Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2000) NOTE: Use nuclease-free water to make all the reagents. Milli-Q water or equivalent grade of ultrapure water can be used for the experiments with RNA. Wear disposable gloves and mask.

Prepare the cell lysate 1. Wash the cells once with 2 ml of CMF-DPBS. 2. Aspirate CMF-DPBS completely, and add 1 ml of Trizol reagent, and incubate for 5 min at room temperature. 3. Collect the cell lysate in 1.5-ml microcentrifuge tube. You can stop the experiment after completing this step. Cell lysates should be stored at −80◦ C.

Purify the RNA 4. Add 200 μl of chloroform to the thawed lysate and mix vigorously by shaking. 5. Centrifuge 5 min at 15,000 × g, room temperature. 6. Transfer the aqueous phase (500 μl) to a new 1.5-ml microcentrifuge tube, add 400 μl of isopropanol, and mix well by inversion for 20 min. 7. Centrifuge tube 5 min at 15,000 × g, room temperature. 8. Remove the isopropanol, add 500 μl of 70% ethanol and centrifuge 5 min at 15,000 × g, room temperature. 9. Remove the ethanol completely and air dry the pellet at room temperature for 2 to 3 min.

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PCR primers and reaction conditions for pluripotent cell marker analysis. Figure 4A.2.3

Generation and Characterization of Human iPS Cells

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Current Protocols in Stem Cell Biology

10. Resuspend the pellet in 26 μl of RNase-free water. You can stop the experiment after completing this step. Purified RNA samples should be stored at −80◦ C.

Remove genomic DNA contamination by DNase treatment 11. Add 3 μl of 10× DNase I buffer and 1 μl of DNase I (from Turbo DNA-free kit) to the RNA sample, mix gently by finger tapping, and incubate for 30 min at 37◦ C. 12. Add 3 μl of DNase Inactivation Reagent (from Turbo DNA-free kit), and mix well. 13. Incubate for 3 min at room temperature with occasional mixing by finger tapping. 14. Centrifuge 3 min at 15,000 × g, room temperature. Transfer the supernatant carefully to a new 1.5-ml microcentrifuge tube.

Determine RNA concentration 15. Use 1 μl of DNase-treated sample to determine concentration of RNA samples by measuring A260 /A280 with an optical spectrometer (e.g., Nanodrop), and adjust concentration of each sample to appropriate one. Samples should be >100 ng/μl RNA for RT-PCR. You can stop the experiment at this step. Purified RNA samples should be stored at −80◦ C.

Perform reverse transcription 16. Prepare 20 μl of reaction mixture by mixing the reagents listed below: 4 μl 5× reverse transcription buffer (from ReverTra Ace kit) 2 μl 10 mM dNTPs (from ReverTra Ace kit) 1 μl ribonuclease inhibitor (from ReverTra Ace kit) 1 μl ReverTra Ace (reverse transcriptase; from ReverTra Ace kit) 1 μl 10 μM oligo dT20 primer (from ReverTra Ace kit) 1 μg DNase-treated total RNA (step 14) Nuclease-free water up to 20 μl. You should prepare reactions containing no reverse transcriptase as negative controls for each sample.

17. Incubate the mixture in thermal cycler at the condition as follows:

60 min at 42◦ C 5 min at 95◦ C Indefinitely at 4◦ C. You can stop the experiment at this step. cDNA samples should be stored at −20◦ C or lower.

Amplify the products by PCR 18. Prepare 25 μl of PCR mixture by mixing the reagents listed below in a 0.2-ml PCR reaction tube: 2.5 μl 10× ExTaq buffer (from ExTaq kit) 2 μl 2.5 mM dNTPs (from ExTaq kit) 0.5 μl 10 μM forward primer (Figure 4A.2.3) 0.5 μl 10 μM reverse primer (Figure 4A.2.3) 0.5 μl 5 U/μl ExTaq DNA polymerase (from ExTaq kit) 1 μl cDNA sample (step 17) 1.25 μl of DMSO (optional, depends on primer sets) Nuclease-free water up to 25 μl.

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19. Carry out PCR according to the conditions listed in Figure 4A.2.3. PCR conditions, particularly the number of cycles, may differ among different thermal cyclers. It is necessary to experiment to find the optimal conditions.

20. After finishing PCR, analyze the by electrophoresis on a 2% agarose gel in 1× TAE buffer using a standard protocol (e.g., Voytas, 2000). SUPPORT PROTOCOL 4

IMMUNOCYTOCHEMISTRY FOR PLURIPOTENT CELL MARKERS The expression of pluripotent stem cells marker can be confirmed not only by RT-PCR (Support Protocol 3) but also by immunocytochemistry. Some surface antigens specifically expressed in pluripotent cells such as SSEAs and TRAs were identified by analyses of human embryonic carcinoma (EC) and ES cells. See Figure 4A.2.2 for examples of immunohistochemistry results.

Materials Human iPS cells (Basic Protocol 1) 6-well plates seeded with mitomycin C–treated feeder cells (Support Protocol 1) hES medium (see recipe) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CMF-DPBS containing 10% (v/v) formalin CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum (or donkey serum), and 0.2% (v/v) Triton X-100 (omit Triton if staining surface antigens) Primary antibodies against desired ES markers (perform all dilutions in CMF-DPBS containing 1% v/v bovine serum albumin): Anti-Nanog goat polyclonal (R&D Systems, cat. no. AF1997; use at 1:20 dilution) Anti-SSEA-1 mouse IgM (Developmental Studies Hybridoma Bank, cat. no. MC480; use at 1:5 dilution) Anti-SSEA-3 rat IgM (Developmental Studies Hybridoma Bank cat. no. MC631; use at 1:5 dilution) Anti-TRA-1-60 mouse IgM (Chemicon, cat. no. MAB4630; use at 1:50 dilution) Anti-TRA-1-81 mouse IgM (Chemicon, cat no. MAB4381; use at 1:50 dilution) Secondary antibody against IgG or IgM of species in which primary antibody was raised, labeled with Alexa Fluor 488 or Alexa Fluor 546; use at 1:500 dilution in CMF-DPBS containing 1% (w/v) bovine serum albumin 10 mg/ml Hoechst 33342 (H3570, Invitrogen) 1. To prepare cells for immunostaining, seed about 100 to 200 clumps of human iPS cells in hES cell medium in each well of a 6-well plate containing mitomycin-treated SNL feeder cells and incubate for 5 to 7 days prior to fixation.

Fix cells and block nonspecific binding 2. Prior to fixation, aspirate the medium, and wash with 2 ml of CMF-DPBS. 3. Remove CMF-DPBS, add 2 ml of CMF-DPBS containing 10% formalin, and fix the cells by incubating for 10 min at room temperature. 4. After fixation, wash the cells once with 2 ml of CMF-DPBS.

Generation and Characterization of Human iPS Cells

5. Aspirate CMF-DPBS and add 2 ml of CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum, and 0.2% (v/v) Triton X-100. Incubate 45 min at room temperature.

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Omit Triton X-100 when staining for surface antigens. Triton X-100 is not necessary for immunostaining of surface antigens. Treatment with Triton X-100 is required only for anti-Nanog antibody. For anti-Nanog antibody, substitute normal donkey serum for normal goat serum because anti-Nanog antibody was raised in goat.

Treat cells with primary and secondary antibodies 6. After blocking procedure, incubate the cells 1 ml of primary antibody at the appropriate dilution in CMF-DPBS containing 1% bovine serum albumin, overnight at 4◦ C. Other antibodies should work. Determine the optimal dilution.

7. Wash the cells three times each for 5 min with CMF-DPBS. 8. Add 1 ml of secondary antibody conjugated with Alexa Fluor 488 or 546 to the sample at the appropriate dilution in CMF-DPBS containing 1% bovine serum albumin supplemented with 1 μg/ml of Hoechst 33342 (added from 10 mg/ml Hoechst stock solution), and incubate for 45 min at room temperature in the dark. 9. Wash out secondary antibody with 2 ml CMF-DPBS three times, each time for 5 min. 10. Observe the cells with a fluorescent microscope equipped with the appropriate filters.

ASSESSING PLURIPOTENCY BY IN VITRO DIFFERENTIATION OF iPS CELLS BY EMBRYOID BODY FORMATION

SUPPORT PROTOCOL 5

Embryoid body formation is one of the easiest procedures for in vitro differentiation of ES cells. This also can be applied for differentiation of iPS cells. Our protocol consists of a primary floating culture for 8 days. After 8 days of floating culture, transfer the cells to gelatin-coated plates to induce further differentiation. After embryoid body formation, differentiation should be confirmed by immunocytochemistry for differentiated markers. Other procedures such as RT-PCR (Support Protocol 3) are also suitable for determination of pluripotency and/or differentiation.

Materials 10 mg/ml HEMA-MMA (see recipe) Growing human iPS cells (Basic Protocols 1 and 2) at 80% to 90% confluency in 60-mm dish Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium containing no bFGF (see recipe) CMF-PBS containing 10% (v/v) formalin (Sigma) CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum (or donkey serum), and 0.2% (v/v) Triton X-100 Primary antibodies against desired ES markers for immunohistochemistry (perform all dilutions in CMF-PBS containing 1% v/v bovine serum albumin): Anti-α-fetoprotein mouse IgG (R&D Systems, cat. no. MAB1368; use at 1:100 dilution) Anti-α-smooth muscle actin mouse IgG (Dako, cat. no. N1584; use at 1:500 dilution) Anti-βIII-tubulin mouse IgG (Chemicon, cat. no. CB412; use at 1:100 dilution) Secondary antibody: anti-mouse IgG labeled with Alexa Fluor (use at 1:500 dilution in CMF-DPBS containing 1% w/v bovine serum albumin) 10 mg/ml Hoechst 33342 solution (Invitrogen)

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100-mm tissue culture dish Sterile disposable cell scraper 15-ml conical centrifuge tubes Gelatin-coated 6-well culture plate (see recipe) Additional reagents and equipment for immunohistochemistry (Support Protocol 4) Establish suspension culture 1. Place 5 ml of 10 mg/ml of HEMA-MMA in a 100-mm dish, and incubate at room temperature in a hood with the dish covered with foil until the solution dries up (3 to 5 days). 2. Wash the iPS cells in 60-mm dish once with 4 ml CMF-DPBS. 3. And add 0.5 ml of CTK solution and return the dish to the 37◦ C incubator. 4. After 5 min incubation, wash twice with 4 ml of CMF-DPBS to remove the CTK solution and detached feeder cells. 5. Add 4 ml hES medium without bFGF to the dish. 6. Detach iPS colonies from the dish by using cell scraper. Collect the cell clumps to a 15-ml conical tube. Do not break up the colonies; larger colonies can form embryoid bodies effectively.

7. Add another 5 ml of hES medium without bFGF and transfer the cell suspension to the HEMA-coated 100-mm dish from step 1. 8. Incubate 2 days in humidified 37◦ C, 5% CO2 incubator. 9. To change the medium, collect the cell suspension into a 15-ml conical tube and let sit it for 5 min at room temperature. 10. Remove the supernatant (∼8 ml) carefully, then add 8 ml of fresh hES medium without bFGF and return the suspension to a HEMA-coated dish prepared as in step 1. Change the medium every other day.

Set up attached culture 11. Collect the iPS cell suspension into a 15-ml conical tube, and let sit for 5 min at room temperature. Remove the supernatant and resuspend the cells in 12 ml of hES medium without bFGF. 12. Transfer 2 ml of cell suspension into wells of a gelatin-coated 6-well culture plate, and incubate at 37◦ C, 5% CO2 . 13. Change the medium every other day. 14. After 8-day attached culture, perform immunocytochemistry for differentiated cell markers (see Support Protocol 4). We routinely observe the expression of α-fetoprotein for endoderm, α–smooth muscle actin for mesoderm, and βIII-tubulin for ectoderm. Other antibodies and markers may also be used for this purpose. SUPPORT PROTOCOL 6 Generation and Characterization of Human iPS Cells

ASSESSING PLURIPOTENCY BY IN VIVO DIFFERENTIATION BY TERATOMA FORMATION Teratoma formation is another well known, important test of pluripotency. In general, mouse ES and iPS cells can produce teratomas easily. However, it is hard to form tumors derived from either human ES or iPS cells by subcutaneously injection into immunodeficient mice, including NOD-SCID mice. Therefore, in this protocol we inject

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stem cells into testes of SCID mice. This change improves the efficiency of tumor formation to more than 80%.

Materials 10 mM Y-27632 (Wako, cat. no. 253-00513) Growing iPS cells (Basic Protocols 1 and 2) at 80% to 90% confluency in 60-mm dish Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium (see recipe) DMEM/F12 medium (e.g., Invitrogen) supplemented with 10 μM Y-27632 1.2% tribromoethanol (Avertin): dissolve 2.5 g tribromoethanol in 5 ml butanol, then add 200 ml distilled water; store at 4◦ C in the dark SCID mice, (7- to 8-weeks, male) 70% ethanol CMF-PBS containing 10% formalin Sterile disposable cell scrapers 15-ml conical centrifuge tubes Centrifuge Hamilton syringe 25-G to 26-G needle (Terumo) Suture needle with thread Additional reagents and equipment for intraperitoneal injection (Donovan and Brown, 2006a) and euthanasia of the mouse (Donovan and Brown, 2006b), paraffin embedding and sectioning of tissue, and hematoxylin/eosin staining of tissue sections (UNIT 2A.5) NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to government regulations for the care and use of laboratory animals.

Prepare cell suspension 1. Add 10 μM Y-27632 (from 10 mM stock) to the medium of a confluent culture of iPS cells, and incubate at 37◦ C for at least 1 hr. Y-27632 is a specific inhibitor for p160-Rho-associated coiled-coil kinase (ROCK).

2. Wash the cells with 4 ml of CMF-DPBS, and add 0.5 ml of CTK solution. Incubate ∼5 min at room temperature. After treatment with Y-27632, the cells may become less detachable. In such cases, you can treat the cells with CTK solution for longer period of time (∼10 min).

3. Wash out CTK solution and detached feeder cells with 4 ml of CMF-DPBS, twice, and add 4 ml of hES medium. 4. Detach iPS cells from the dish with a cell scraper, and break the colonies into small clumps by pipetting up and down several times. 5. Collect the cell suspension to a 15-ml conical tube, and centrifuge 5 min at 200 × g, room temperature. 6. Aspirate the supernatant, and resuspend cells in 300 to 500 μl of DMEM/F12 supplemented with 10 μM of Y-27632. Manipulation of Potency

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Inject cells 7. Inject 0.8 ml of 1.2% tribromoethanol solution intraperitoneally (Donovan and Brown, 2006a) into SCID mouse (0.12 ml for 10 g weight). 8. Wash the lower abdominal/groin area with 70% ethanol. 9. Dissect out the testes and remove from the body. Dissect the lower abdominal/groin area and withdraw the inguinal canal and then the testes. Leave spermatic cord intact. 10. Inject 30 μl of iPS cell suspension into a testes, under the capsule, using a Hamilton syringe and a 25-G to 26-G needle, as gently as possibly. 11. Return the testes to the original interperitoneal location, and close the incision with stitches. Return mouse to colony within 2 hr. 12. About 3 months later, observe the mice for teratoma formation (Fig. 4A.2.2K). Mice may appear to be pregnant, indicating the presence of a teratoma.

Dissect the tumors 13. Euthanize mice (Donovan and Brown, 2006b) bearing teratomas and dissect out the tumors. 14. Fix the tumors in ∼50 ml of CMF-DPBS containing 10% formalin and incubate overnight at room temperature with agitation. 15. After fixation, embed the tumor in paraffin. 16. Slice the tumor into 4- to 5-μm sections and mount on slides. 17. Stain the sections with hematoxylin and eosin using a standard protocol (e.g., UNIT 2A.5). 18. Examine the entire set of sections for a tumor, scoring for the presence of derivatives of all three germ layers such as cartilage, pigmented epithelium, and gut-like epithelium (see Fig. 4A.2.2M). If the tumor contains derivatives of all three germ layers, the iPS cell line is pluripotent.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

10% FBS medium DMEM (e.g., Invitrogen) containing: 10% fetal bovine serum (FBS) 50 U/ml penicillin 50 μg/ml streptomycin To prepare 500 ml of 10% FBS medium, mix 50 ml FBS and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week.

Generation and Characterization of Human iPS Cells

For Plat-E cells (see Support Protocol 2), add 1 μl of 10 mg/ml puromycin stock (see recipe) and 10 μl of 10 mg/ml blasticidin S stock (see recipe) to 10 ml of 10% FBS medium.

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293FT medium DMEM (e.g., Invitrogen) containing: 10% fetal bovine serum (FBS) 2 mM L-glutamine 1 × 10−4 M nonessential amino acids 1 mM sodium pyruvate 50 U penicillin 50 μg/ml streptomycin 0.5 mg/ml G418 To prepare 500 ml of the medium, mix 50 ml of FBS, 5 ml of 200 mM (100×) L-glutamine, 5 ml 100× nonessential amino acids, 5 ml of 100 mM sodium pyruvate, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week. Just before use, add 100 μl of 50 mg/ml G418 to 10 ml 293FT medium.

Blasticidin S stock solution Dissolve blasticidin S hydrochloride (Funakoshi Chemical Company; http://www. funakoshi.co.jp) in distilled water at 10 mg/ml and sterilize through a 0.22-μm filter. Aliquot and store at −20◦ C.

CTK solution 5 ml 2.5% (w/v) trypsin 5 ml 1 mg/ml collagenase IV 0.5 ml 0.1 M CaCl2 10 ml Knockout Serum Replacement (KSR; Invitrogen) 30 ml distilled water Store up to 1 month at −20◦ C Do not repeat freeze/thaw cycles DAP213 solution To 5.37 ml hES medium (see recipe) add: 1.43 ml DMSO 1 ml 10 M acetamide 2.2 ml of propylene glycol Store up to 1 month at −80◦ C Gelatin coating of culture vessels Dissolve 1 g of gelatin powder (Sigma, cat. no. G-1890) in 100 ml of distilled water, autoclave, and store at 4◦ C as the 10× gelatin stock solution. To prepare 0.1% (1×) gelatin solution, thaw the 10× gelatin stock in a microwave and/or autoclave, then add 50 ml of the 10× solution to 450 ml of distilled water. Filter the solution with a 0.22-μm filter unit and store at 4◦ C. To coat culture dishes, add appropriate volume of 0.1% (1×) gelatin solution to cover the entire area of the dish bottom. For example, 1, 3, or 5 ml of gelatin solution is used for a 35-, 60-, or 100-mm dish, respectively. Incubate the dishes for at least 30 min at 37◦ C in a sterile environment. Before using, aspirate off the excess gelatin solution. Gelatin stock is prepared as 10× concentration (1% w/v) stocks.

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1 × 10−4 M nonessential amino acids 1 × 10−4 M 2-mercaptoethanol 50 U penicillin 50 μg/ml streptomycin To prepare 500 ml of the medium, mix 100 ml KSR, 5 ml of 200 mM (100×) L-glutamine, 5 ml 100× nonessential amino acids, 1 ml 2-mercaptoethanol, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM/F12. Add 200 μl of 10 μg/ml bFGF into 500 ml of the medium just before use. For differentiation experiments (e.g., Support Protocol 5), do not add bFGF. Store at 4◦ C up to 1 week. All abovementioned components are available from Invitrogen. Primate ES cell medium from ReproCELL (http://www.reprocell.net/) may be used as an alternative.

Mitomycin C, 0.4 mg/ml Dissolve 2 mg of mitomycin C (Kyowa Hakko Kirin; http://www.kyowa-kirin.co.jp/ english/) in 5 ml of CMF-DPBS (Nacalai Tesque, cat. no. 14249-95). Store up to 1 month at −20◦ C in the dark. CAUTION: Because of its toxicity, the solution must be treated exclusively in a safety cabinet with gloves and lab coat and disposed of in accordance with the rules each institution stipulates.

Poly(hydroxyethyl methacrylate-co-methyl methacrylate; HEMA-MMA), 10 mg/ml Add 0.3 g of HEMA-MMA (Sigma, cat. no. P-3932) to a tube containing 30 ml ethanol. Incubate at 37◦ C overnight with agitation. Prepare fresh for each experiment.

Puromycin Dissolve puromycin (Sigma, cat. no. P-8833) in distilled water at 10 mg/ml and sterilize through a 0.22-μm filter. Divide into aliquots and store up to 1 month at −20◦ C.

SNL medium DMEM (e.g., Invitrogen) containing: 7% fetal bovine serum (FBS) 2 mM L-glutamine 50 U penicillin 50 μg/ml streptomycin To prepare 500 ml of the medium, mix 35 ml FBS, 5 ml 200 mM (100×) Lglutamine, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week. This medium is used for fibroblasts and PLAT-E cells.

COMMENTARY Background Information

Generation and Characterization of Human iPS Cells

Although it is commonly known that nuclei of differentiated cells can be reprogrammed back to embryonic states by means of nuclear transfer into oocytes or fusion with ES cells, the mechanism of inducing nuclear reprogramming has yet to be revealed. The fact that

somatic cells can be reprogrammed by fusion with ES cells implies that ES cells contain factors that can induce reprogramming. We hypothesized that factors which play important roles in ES cells also play pivotal roles in induction of nuclear reprogramming. Pluripotency and tumor-like proliferation are

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the most exquisite properties of ES cells. Three transcription factors—Oct3/4, Sox2, and Nanog—have been found to be essential in the maintenance of pluripotency in both early embryos and ES cells. While a handful of laboratories have demonstrated that several tumor-related gene products, such as ERas, c-myc, and Stat3, contribute to long-term maintenance of ES cells in culture, we have identified several genes that are specifically expressed in ES cells by analyzing expressed sequence tag (EST) databases. After selecting the most promising 24 gene products as candidates for potential factors that could induce reprogramming, we narrowed these to four transcription factors (Oct3/4, Sox2, Klf4, and c-myc) that have been shown to convert fibroblasts back to pluripotent state. The identification of these factors was an important breakthrough that has revealed a mechanism of nuclear reprogramming and let us create pluripotent cells directly from skin biopsy specimens. One year later, other groups succeeded in generating iPS cells from human somatic cells. Recently, two research groups have reported that various disease-specific iPS cells from a patient’s own somatic cells have been successfully reprogrammed (Dimos et al., 2008; Park et al., 2008b). Now, iPS cell technology can be used in conjunction with or in place of ES cell technology to shed light on understanding pathogens, in drug discovery, and most of all, to develop regenerative medicine applications. Encouraging broad use of iPS cell technology will facilitate the development of practical applications. These protocols should provide guidance to scientists who share our objectives.

Troubleshooting In some cases, lentivirus is toxic to fibroblasts. Depending on the different cell lines, lentiviral transduction may lead to loss or growth arrest of fibroblasts due to their sensitivity to the virus. As some fibroblasts are more vulnerable to lentivirus than common cells, they should be treated with a double dilution of the virus-containing supernatant in fresh medium or by shortening the exposure time from overnight to 5 hr. For our purposes, the expression of mouse Slc7a1 gene is sufficient for generation of iPS cells despite lower infection efficiency. When no ES-like (iPS) colonies appear in fibroblast cultures after introduction of the four factors, the following causes should be considered. First, the titer of retrovirus may be too

low. Transduction efficiencies of retroviruses for reprogramming factors are critical for iPS cell colony formation as described above. The retrovirus must be prepared fresh for every experiment. Do not use frozen stock retroviruses because freezing causes reduction of the titer. Growth properties of the fibroblasts are also important for iPS cell generation. Efficiency of retroviral transduction is markedly reduced when senescent fibroblasts are used for transduction. We strongly recommend banking stocks of fibroblasts at early passages and using fresh fibroblasts of early passage for iPS cell production. The number of cells that are plated onto SNL feeder cells after retroviral transduction is important. Overgrowth of fibroblasts might make cells peel off from the edge of the dish like a sheet, inhibiting formation of iPS cell colonies. Although this may be overcome by reducing the cell number, too small a number of cells could lead to no colony appearance. The optimal conditions differ for each individual cell type. We recommend that you seed at least in two or three different dishes with different densities when first plating the transduced cells. In addition, the qualities of feeder cells are crucial not only for generation of iPS cells, but also for maintenance of them. If feeder cells are too old, cells may peel off the substrate during the reprogramming or maintenance. SNL feeder cells more than 3 days after mitomycin C treatment cannot survive the stimulation by bFGF in hES medium (as bFGF may have a toxic effect on older feeder cells). It is recommended that SNL feeder cells be used within 3 days after inactivation. Some problems may arise after iPS cells are generated. For example, iPS cells can change characteristics and potential, depending on the line, with long-term culture. Human iPS cells, like human ES cells, may become adapted in a long-term culture. We recommend that large amounts of iPS cell stocks be frozen at early passages to support long-term experimentation. iPS cells are relatively unstable during early passage period so that spontaneous differentiation in daily culture may also happen. When the number of differentiated colonies increases, select undifferentiated colonies and transfer them by aspiration to a new dish of SNL feeder cells. After this procedure is repeated two to three times, the majority of the dish will consist of undifferentiated colonies. In addition, the qualities of feeder cells, such as density and freshness, are also important.

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Anticipated Results The efficiency of lentiviral transduction to fibroblasts should be >90%. You can estimate the efficiency of infection with the GFPencoding lentivirus. On the other hand, because retroviruses can be transfected only into dividing cells, the transduction efficiency may stay at ∼30% to 60%. From 10 days to 2 weeks after retroviral transduction, some granule colonies usually appear (Fig. 4A.2.2, panel A). However, these colonies are not iPS cells. Generally, clearedged colonies are produced at 3 weeks post transduction (panel B). They can be expanded after being picked up and transferred to another plate. Established iPS cells show hESlike morphologies on feeder cells (panel C). When the cells reach this stage, you should passage once a week. The expression of markers in pluripotent stem cells can be detected in iPS cells as similar level to ES cells. iPS cells typically express SSEA3 (panel D), TRA-1-60 (panel E), TRA-1-81 (panel F), and Nanog (panel G), but not SSEA1 (panel H). Differentiation potentials of iPS cells can be determined easily by embryoid body formation. After a 16-day induction, the expression of differentiation markers such as αfetoprotein (panel I), α-smooth muscle actin (panel J) and βIII-tubulin (panel K) can be confirmed by immunocytochemistry. Another assay for determination of pluripotency, teratoma formation, is also important. Generally, around 3 months after injection of iPS cells into the testes of SCID mice, the mice may appear to be pregnant (panel L). In some cases, black-colored pigment cells can be observed in dissected tumors by the naked eye (panel M). Staining of tumors with hematoxylin and eosin may show that many types of all three germ layers exist in the teratoma if parental iPS cells are pluripotent (panel N). Treatment of human iPS cells with Y-27632, which is an inhibitor for p160-Rhoassociated coiled-coil kinase (ROCK), before harvesting, may improve the survival rate. If you have trouble with frail viability of iPS cells, you can treat the cells at least an hour before harvesting.

Time Considerations

Generation and Characterization of Human iPS Cells

It takes 1 week to successfully transduce the fibroblasts with the lentiviral vector and to verify transduction by microscopic examination or flow cytometry. Then it requires an additional 5 days to prepare the retrovirus vectors

and transduce the fibroblasts. Once plated on SNL feeder cells, it takes ∼3 weeks for iPS cell colonies to appear and additional time for them to grow to a size where they can be passaged. iPS cells are fed every other day and passaged once a week. Overall, it takes over 3 months to establish an iPS cell line.

Acknowledgements We thank Dr. Tetsuya Ishii, Kanon Takeda, and Yuko Shimazu for reading the manuscript, and Tomoko Ichisaka and Noriko Tsubooka and other members of Yamanaka laboratory for valuable discussion and support. Thanks too to Rie Kato, Ryoko Iyama, Eri Nishikawa, Noriyo Maruhashi and the member of CiRA for administrative support. We are also grateful to Drs. Toshio Kitamura for retroviral system, Peter Andrews for antibodies, and Yoshiki Sasai for instruction on teratoma experiments.

Literature Cited Blelloch, R., Venere, M., Yen, J., and RamalheSantos, M. 2007. Generation of induced pluripotent stem cells in the absence of drug selection Cell Stem Cell 1:245-247. Dimos, J.T., Rodolfa, K.T., Niakan, K.K., Weisenthal, L.M., Mitsumoto, H., Chung, W., Croft, G.F., Saphier, G., Leibel, R., Goland, R., Wichterle, K., Henderson, C.E., and Eggan, K. 2008. Induced pluripotent stem cells generated from patients with ALS can be differentiated into motor neurons. Science 321:1218-1221. Donovan, J. and Brown, P. 2006a. Parenteral injections. Curr. Protoc. Immunol. 73:1.6.1-1.6.10. Donovan, J. and Brown, P. 2006b. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Fujioka, T., Yasuchika, K., Nakamura, Y., Nakatsuji, N., and Suemari, H. 2004. A simple and efficient cryopreservation method for primate embryonic stem cells. Int. J. Dev. Biol. 48:1149-1154. Lowry, WE., Richter, L., Yachenko, R., Ryle, A.D., Tchieu, J., Sridharan, R., Clark, A.T., and Plath, K. 2008. Generation of human induced pluripotent stem cells from dermal fibroblasts. Proc. Natl. Acad. Sci. U.S.A. 105:2883-2888. Maherali, N., Sridharan, R., Xie, W., Utikal, J., Eminli, S., Arnold, K., Stadtfeld, M., Yacheehkes, R., Tchieu, J., Jaenisch, R., Plath, K., and Hochedinger, K. 2007. Directly reprogrammed fibroblasts show global epigenetic remodeling and widespread tissue contribution. Cell Stem Cell 1:55-70. Masaki, H., Ishikawa, T., Takahashi, S., Okumura, M., Sakai, N., Haga, M., Kominami, K., Migita, H., McDonald, F., Shimada, F., and Sakurada, K. 2008. Heterogeneity of pluripotent marker gene expression in colonies generated in human iPS cell induction culture. Stem Cell Res. In press.

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McMahon, A.P. and Bradley, A. 1990. The Wnt-1 (int-1) proto-oncogene is required for development of a large region of the mouse brain. Cell 62:1073-1085. Meissner, A., Wernig, M., and Jaenisch, R. 2007. Direct reprogramming of genetically unmodified fibroblasts into pluripotent stem cells. Nat. Biotech. 25:1177-1181. Morita, S., Kojima, T., and Kitamura, T. 2000. Plat-E: An efficient and stable system for transient packaging of retroviruses. Gene Ther. 7:1063-1066. Nakagawa, M., Koyanagi, M., Tanabe, K., Takahashi, K., Ishisaka, T., Aoi, T., Okita, K., Mochiduki, Y., Takizawa, N., and Yamanaka, S. 2008. Generation of induced pluripotent stem cells without Myc from mouse and human fibroblasts. Nat. Biotech. 26:101-106. Okita, K., Ishisaka, T., and Yamanaka, S. 2007. Generation of germline-competent induced pluripotent stem cells. Nature 448:313-317. Park, I.H., Zhao, R., West, J.A., Yabuuchi, A., Huo, H., Ince, T.A., Leroy, P.H., Lensch, M.W., and Daley, G.O. 2008a. Reprogramming of human somatic cells to pluripotency with defined factors. Nature 451:141-146. Park, I.H., Arora, N., Huo, H., Maheraum, N., Ahfeldt, T., Shimamuki, N., Lensch, M.W., Cowan, C., Hochedinger, K., and Daley, G.O. 2008b. Disease-specific induced pluripotent stem cells. Cell 134:877-886. Takahashi, K. and Yamanaka, S. 2006. Induction of pluripotent stem cells from embryonic and

adult fibroblast cultures by defined factors. Cell 126:663-676. Takahashi, K., Tanabe, K., Ohnuki, M., Narita, T., Tomoda, K., and Yamanaka, S. 2007a. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861-872. Takahashi, K., Okita, K., Nakagawa, M., and Yamanaka, S. 2007b. Induction of pluripotent stem cells from fibroblast cultures. Nat. Protoc. 2:3081-3089. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Watanabe, K., Ueno, M., Kamiya, D., Nishiyama, A., Matsumura, M., Wataya, T., Takahashim, J.B., Nishikawa, S., Nishikawa, S., Miguruma, K., and Sasai, Y. 2007. A ROCK inhibitor permits survival of dissociated human embryonic stem cells. Nat. Biotech. 25:681-686. Wernig, M., Meissner, A., Foreman, R., Bambrook, T., Ku, M., Hochedinger, K., Bernstein, R.E., and Jaenisch, R. 2007. In vitro reprogramming of fibroblasts into a pluripotent ES-cell-like state. Nature 448:318-324. Yamanaka, S. 2007. Strategies and new developments in the generation of patient-specific pluripotent stem cells. Cell Stem Cell 1:39-49. Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S., Nie, J., Jonsdottir, G.A., Ruotti, V., Stewart, R., Slukvin, I.I., and Thomson, J.A. 2007. Induced pluripotent stem cell lines derived from human somatic cells. Science 318:1917-1920.

Manipulation of Potency

4A.2.25 Current Protocols in Stem Cell Biology

Supplement 9

Heterokaryon-Based Reprogramming for Pluripotency

UNIT 4B.1

Carlos Filipe Pereira1 and Amanda G. Fisher1 1

Imperial College School of Medicine, Hammersmith Hospital, London, United Kingdom

ABSTRACT Embryonic stem (ES) cells have the ability to self-renew, execute multiple lineage paths, and dominantly reprogram differentiated cells upon cell fusion. Here, we describe an approach that reprograms human B lymphocytes toward pluripotency by generating inter-species heterokaryons with mouse ES cells. This induces a human ES-specific gene expression profile, in which the extent and the rapidity of conversion allows us to compare the capacity of different mouse ES cell lines to dominantly induce pluripotency. This approach, coupled with pharmacological inhibition, gene knock-out, or knockdown permits factors that are required to directly induce reprogramming to be defined individually, as well as in combination. Experimental heterokaryons provide a simple and tractable approach to address the mechanisms underlying direct reprogramming to pluripotency. The procedure requires 5 days to complete. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 9:4B.1.1-4B.1.14.  Keywords: reprogramming r embryonic stem (ES) cell r pluripotency r cell fusion r heterokaryon

INTRODUCTION Epigenetic reprogramming can be achieved in different ways including nuclear transfer or the forced expression of transcription factors to induce pluripotency (Hochedlinger and Jaenisch, 2006; Yamanaka, 2007). However, the low frequency and/or the long period of time required for inducing pluripotency (Stadtfeld et al., 2008) hinders a systematic appraisal of the mechanisms underlying direct reprogramming to pluripotency. Reprogramming can also be achieved by cellular fusion. Spontaneous and experimental cell fusion of differentiated cells with pluripotent cells induces the expression of pluripotencyassociated markers in the hybrid cells (Tada et al., 2001; Cowan et al., 2005; Silva et al., 2006). Here, we describe a simple protocol for generating heterokaryons (cells in which parental nuclei share the same cytoplasm but remain spatially discrete), in which the nuclear events that occur in the donor and recipient nucleus can be discerned using species-specific reagents (Terranova et al., 2006; Pereira et al., 2008). In addition to inducing pluripotency-associated gene expression by somatic cells, this procedure allows an analysis of epigenetic changes within the reprogrammed nucleus over time, including nuclear re-organization, DNA/chromatin modifications, and the requirement of dominant trans-acting factors (Pereira et al., 2008). This unit describes a protocol for the reprogramming of human somatic cells by the formation of heterokaryons with mouse ES cells. The unit begins with the protocol for heterokaryon formation and gene expression analysis by quantitative RT-PCR (Basic Protocol), and it is followed by an alternative method (Alternate Protocol) that allows for the selection of heterokaryons without pre-labeling and fluorescence-activated cell sorting (FACS).

Manipulation of Potency Current Protocols in Stem Cell Biology 4B.1.1-4B.1.14 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc04b01s9 C 2009 John Wiley & Sons, Inc. Copyright 

4B.1.1 Supplement 9

BASIC PROTOCOL

GENERATION OF INTER-SPECIES HETEROKARYONS BETWEEN HUMAN B LYMPHOCYTES AND MOUSE ES CELLS This protocol is used for the formation of heterokaryons between human somatic cells and mouse ES cells. Human B cells are fused with mouse ES cells using polyethylene glycol (PEG), and the reprogramming of human somatic cells is monitored by human gene–specific quantitative RT-PCR (Fig. 4B.1.1A). Inter-species heterokaryons can be generated by cell fusion of adherent ES cells and lymphocytes (non-adherent cells). The resulting heterokaryons will attach to gelatin-coated dishes or to irradiated mouse embryonic fibroblast (MEF) feeder layers. NOTE: The quality of donor and recipient cells is essential for reprogramming experiments. We suggest that several ES cell lines/clones that are known to give germ-line transmission (donor) are initially tested in the reprogramming assay. In addition, different somatic cells (recipient) may also be more or less prone to successful reprogramming. NOTE: In addition to gene expression, inter-species heterokaryons can be generated in parallel for other applications (Fig. 4B.1.1A), including bisulfite genomic sequencing (BGS), immunofluorescence (IF; Fig. 4B.1.1C), fluorescence in situ hybridization (FISH), and FACS analysis (Pereira et al., 2008).

Materials

HeterokaryonBased Reprogramming for Pluripotency

Epstein-Barr virus (EBV)–transformed human B cell clones Human B cell medium (hB cell medium; see recipe) Mouse ES/heterokaryon medium (see recipe) Mouse ES cells cultured on gelatin-coated dishes or using mouse embryonic fibroblasts (MEFs) as feeder layers (UNIT 1C.4) Mitotically inactivated MEFs (UNIT 1C.3) 50% (w/v) Polyethylene glycol (PEG) 1500 in 75 mM HEPES, pH 8.0 (Roche, cat. no. 10783641001) Knockout (KO) Dulbecco’s Modified Eagle’s Medium (KO-DMEM; Invitrogen, cat. no. 10829-018) Calcium- and magnesium-free phosphate-buffered saline without (CMF-PBS; Invitrogen, cat. no. 14190-094) 0.05% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25300-054) Vybrant multicolor cell-labeling kit (Molecular Probes, cat. no. V22889) containing: 1,1 -dioctadecyl-3,3,3 ,3 -tetramethylindodicarbocyanine (DiD) cell labeling solution 1,1 -dioctadecyl-3,3,3 ,3 -tetramethylindocarbocyanine perchlorate (DiI) cell labeling solution Leukemia inhibitory factor (LIF; Esgro, Chemicon/Millipore, cat. no. ESG1107) Liquid N2 Mouse monoclonal anti–human LaminA/C (Vector, cat. no. VP-L550), optional Vectashield with DAPI (0.1 μg/ml; Vector, cat. no. H-1200), optional Alexa Fluor 568 phalloidin (Molecular Probes, cat. no. A12380), optional FACS buffer (see recipe) RNA-BEE RNA isolation solvent (AMS Biotechnology, cat. no. CS-501B) DEPC-treated water (Ambion, cat. no. 9915G) TURBO DNA-free kit (Ambion, cat. no. 1907) Superscript first-strand synthesis system (Invitrogen, cat. no. 18080-085) containing: First-strand buffer Superscript III 10 mM dNTP mix (Invitrogen, cat. no. 18427-013)

4B.1.2 Supplement 9

Current Protocols in Stem Cell Biology

Oligo (dT)12-18 primers (Invitrogen, cat. no. 18418-012) RNaseOUT (Invitrogen, cat. no. 10777-019) SYBR Green Master Mix (Qiagen, cat. no. 204145) Human gene-specific primers (see Table 4B.1.1) 175-cm2 tissue culture flasks Gelatin-coated 90-mm tissue culture dishes (see recipe) 37◦ C water bath 37◦ C, 5% CO2 incubator 10-ml pipet 50- and 15-ml conical tubes (BD Falcon) Hemacytometer Conical 30-ml universal tubes (Sterilin, cat. no. 128A) Pasteur pipets 70-μm cell strainer (BD Falcon, cat. no. 352350) 5-ml polystyrene round-bottom tubes (BD Falcon, cat. no. 352054) FACS DiVa cell sorter (Becton Dickinson) or similar Nanodrop ND-1000 spectrophotometer Dyad DNA engine 96-well plates for PCR (Bio-Rad, cat. no. MLL-9651) Real-time PCR engine (MJ research Chromo4) Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1c.3), growing mouse ES cell culture on feeders (UNIT 1C.4), for mitotically inactive mouse embryonic fibroblasts (UNIT 1C.3), for RNA extraction (Kingston et al., 1996), and for RT-PCR (Giulietti et al., 2001) NOTE: The following procedures (steps 1 to 36) are performed in a Class II biological hazard flow hood or a laminar-flow hood. All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly.

Label mouse ES cells and human B lymphocytes 1. Grow 1 × 108 EBV-hB lymphocytes in suspension using 175-cm2 tissue culture flasks. Plate 2 × 107 EBV-hB cells in 200 ml of hB cell medium. Change hB cell medium every 48 hr by centrifuging 5 min at 200 × g, room temperature. 2. In parallel, grow 1 × 108 mouse ES cells on five 90-mm gelatin-coated dishes or on irradiated MEFs (according to the requirements of the specific ES cell line; UNIT 1C.4). Plate 3 × 106 ES cells per 90-mm dish in 15 ml of ES medium. Change medium the following day and collect cells after 48 hr. We recommend to start with a large number of cells since a good fusion efficiency will be ∼10% (Fig. 4B.1.1B). For a kinetic experiment of three time points, 1 × 108 cells of each cell-type are required.

3. Warm the following in a 37◦ C water bath: PEG solution, serum-free KO-DMEM, complete ES/heterokaryon medium, CMF-PBS, and 0.05% trypsin/EDTA. 4. Collect ES cells by adding 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes and incubate at 37◦ C, 5% CO2 for 5 min. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate trypsin. Dissociate ES cell colonies and disrupt cell clumps to single-cells by pipetting up and down. It is important to get a single-cell suspension of ES cells for cell fusion. Clumps of cells will favor ES × ES cell fusions rather than hB × ES fusions. If required, an additional step of resuspension in trypsin before inactivation with medium may be included.

5. Collect both cell types in 50-ml conical tubes and centrifuge 5 min at 200 × g, room temperature. Current Protocols in Stem Cell Biology

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4B.1.3 Supplement 9

A

hB cells DiI+

PEG reprogramming heterokaryon DiI+ DiD+

heterokaryon DiI+ DiD+ mES cells DiD+

time after day 0 cell fusion

day 1

day 2

day 3

FACS sorting of heterokaryons store pellets at -80 C Other applications: IF, FACS, FISH, BGS, etc.

RNA extraction and cDNA synthesis human gene-specific qRT-PCR analysis

C

B human B (hB)

hB + mES

PEG fused (d0) hB:mES 10:1 2.5%

hB:mES 5:1 3.6%

hB:mES 1:1 11.5%

DiD

fluorescence intensity

mouse ES (mES)

DiI

fluorescence intensity

Figure 4B.1.1 (legend at right)

HeterokaryonBased Reprogramming for Pluripotency

4B.1.4 Supplement 9

Current Protocols in Stem Cell Biology

Table 4B.1.1 Human Gene-Specific Primers for qRT-PCR Analysisa

Species/Gene hGapdh

NM 002046

hHprt

NM 000194

hOct4 hNanog hCripto hDnmt3b hTert hTle1 hSox2 hRex1 hCD37 hCD19 hCD45

Sequence 5 -3

Accession number

NM 002701 NM 024865 NM 003212 NM 006892 NM 198253 NM 005077 NM 003106 NM 174900 NM 001774 NM 001770 NM 002838

s

TCTGCTCCTCCTGTTCGACA

as

AAAAGCAGCCCTGGTGACC

s

TCCTTGGTCAGGCAGTATAATCC

as

GTCAAGGGCATATCCTACAACAAA

s

TCGAGAACCGAGTGAGAGGC

as

CACACTCGGACCACATCCTTC

s

CCAACATCCTGAACCTCAGCTAC

as

GCCTTCTGCGTCACACCATT

s

AGAAGTGTTCCCTGTGTAAATGCTG

as

CACGAGGTGCTCATCCATCA

s

GTCAAGCTACACACAGGACTTGACAG

as

AGTTCGGACAGCTGGGCTTT

s

GCCAGCATCATCAAACCCC

as

CTGTCAAGGTAGAGACGTGGCTC

s

TGTCTCCCAGCTCGACTGTCT

as

AAGTACTGGCTTCCCCTCCC

s

CACACTGCCCCTCTCACACAT

as

CATTTCCCTCGTTTTTCTTTGAA

s

GCGTACGCAAATTAAAGTCCAGA

as

CAGCATCCTAAACAGCTCGCAGAAT

s

GTGGCTGCACAACAACCTTATTT

as

GCCTAACGGTATCGAGCGAG

s

GCTCAAGACGCTGGAAAGTATTATT

as

GATAAGCCAAAGTCACAGCTGAGA

s

CCCCATGAACGTTACCATTTG

as

GATAGTCTCCATTGTGAAAATAGGCC

a Adapted from Pereira et al., 2008.

Figure 4B.1.1 (at left) Formation of inter-species heterokaryons between human lymphocytes and mouse ES cells. (A) Shows the experimental strategy used for the generation and analysis of inter-species heterokaryons. Human B lymphocytes (hB) and mouse embryonic stem cells (mES) are respectively labeled with the cell membrane dyes DiI and DiD and fused in the presence of polyethylene glycol (PEG). The resulting heterokaryons (cells in which parental nuclei share the same cytoplasm but remain discrete) are cultured under conditions that promote mES selfrenewal and FACS sorted 1, 2, and 3 days after cell fusion. Total RNA is extracted and reverse transcribed for species-specific qRT-PCR analysis. In parallel, heterokaryons may be generated for other applications including immunofluorescence (IF), FACS analysis, fluorescence in situ hybridization (FISH), and bisulfite genomic sequencing (BGS; Pereira et al., 2008). (B) Fusion efficiency is assessed by FACS analysis as the percentage of double-labeled cells (upper-right quadrant of dot plots). Cell fusion with different hB:mES ratios is shown. (C) hB-derived nuclei are distinguished from mouse nuclei by IF on the basis of DAPI (blue) and human Lamin A/C staining (green). A confocal picture of a heterokaryon [one mouse (with DAPI intense foci, blue) and one human nucleus (hLamin A/C positive, green)] 2 days after cell fusion is shown. Actin staining with phalloidin (red) delineates individual cells.

Manipulation of Potency

4B.1.5 Current Protocols in Stem Cell Biology

Supplement 9

6. Wash both cell types once in 50 ml CMF-PBS and count cells with a hemacytometer (UNIT 1C.3). 7. Resuspend 1 × 108 of each cell type in 20 ml of prewarmed serum-free KO-DMEM. 8. Respectively label ES cells with 50 μl of Vybrant 1,1 -dioctadecyl-3,3,3 ,3 tetramethylindodicarbocyanine (DiD) and hB lymphocytes with 50 μl of 1,1 dioctadecyl-3,3,3 ,3 -tetramethylindocarbocyanine perchlorate (DiI) cell labeling solutions. Incubate for 15 min in a 37◦ C water bath with occasional mixing. Time and dye concentration required for homogeneous labeling depends on the cell type used and has to be optimized. Cells can be analyzed by flow cytometry using the FL-2 (DiI) and FL-4 (DiD) channels (Fig. 4B.1.1B). In our hands, ES cells and lymphocytes can be homogeneously labeled for 15 min at 37◦ C without causing major cell death.

9. Centrifuge cells 5 min at 200 × g, room temperature, and aspirate supernatant to remove the dye. 10. Resuspend cell pellets in 25 ml of prewarmed complete ES/heterokaryon medium for recovery.

Fuse the cells 11. Centrifuge cells 5 min at 200 × g, room temperature. Remove supernatant and resuspend both cell types in 15 ml of prewarmed CMF-PBS. 12. Mix lymphocytes and ES cells in a conical 30-ml universal tube, ratio 1:1. Varying the cell ratio can have a big impact on cell fusion efficiency (Fig. 4B.1.1B). This is dependent on the physical features of cell types used and has to be optimized. In our hands the optimal lymphocyte:ES ratio is 1:1.

13. Centrifuge cell mixture 5 min at 200 × g, room temperature. With a Pasteur pipet, remove the supernatant completely, including drops on the plastic surface. Complete removal of the supernatant is essential to avoid dilution of PEG.

14. Disrupt the pellet by gently tapping the bottom of the tube. 15. Place the cells in a 37◦ C water bath for 2 min. The optimal temperature for cell fusion is 37◦ C. We recommend that both cell pellet/tube and PEG are prewarmed at 37◦ C for cell fusion.

16. Add 1 ml of PEG dropwise over a 60-sec period. Rotate and rock the tube slowly but continuously during this time to promote cell contact. The quality of PEG and pH of the solution is very important for successful cell fusion. The 50% PEG 1500 solution is commercially available (see Materials section) but also can be manually prepared.

17. Place the tube in a 37◦ C water bath and gently continue mixing the cells in PEG for an additional 90 sec. Exact timing in PEG is vital for the outcome.

18. Add 1 ml of serum-free KO-DMEM (prewarmed at 37◦ C) dropwise to the fusion mixture, rotating and rocking the tube, over a 60-sec period. 19. Add 3 ml of KO-DMEM over a 2-min period, continuously mixing the cells. HeterokaryonBased Reprogramming for Pluripotency

20. Slowly add 10 ml of KO-DMEM. The sequence described above is designed to add KO-DMEM slowly to the mixture. The objective is to dilute PEG but to avoid osmotic shock and cell death.

4B.1.6 Supplement 9

Current Protocols in Stem Cell Biology

21. Mix by slowly inverting the tube 4 to 6 times. Small clusters of cells induced by PEG treatment can be observed at this point.

22. Incubate 3 min at 37◦ C.

Plate the heterokaryons 23. Centrifuge fusion products 5 min at 250 × g, 37◦ C. 24. With a Pasteur pipet, aspirate the supernatant. Carefully add 10 ml of prewarmed ES/heterokaryon medium plus 1000 U/ml of LIF, without disrupting the pellet. 25. Incubate the pellet of cells in medium 3 min at 37◦ C. 26. Carefully resuspend cell fusions in the required amount of ES/heterokaryon medium plus 1000 U/ml of LIF. Usually, 15 ml per 90-mm dish is sufficient. If required, fusion efficiency can be assessed at this point directly by FACS analysis (Fig. 4B.1.1B).

27. Remove a small aliquot of cells (usually 1/50 of total) and transfer to a 15-ml centrifuge tube. Centrifuge 5 min at 200 × g, room temperature. 28. Wash the aliquot in 15 ml CMF-PBS and aspirate the supernatant. Snap freeze the pellet in liquid N2 (day 0) and store at −80◦ C. 29. Plate cells in 90-mm gelatin-coated dishes at ∼0.5 × 106 cells/cm2 with 15 ml ES/heterokaryon medium plus 1000 U/ml of LIF. Swirl the dish to ensure distribution over the entire plate surface. 30. Incubate dishes overnight at 37 ◦ C, 5% CO2 . Heterokaryons will attach to the dish.

31. Culture heterokaryons in the same conditions used for mouse ES cells. Change medium daily (the majority of unfused lymphocytes will be removed when changing medium). Heterokaryons can be sorted before or after culturing. However, we recommend sorting after culturing as it results in better yield and decreased cell death. Heterokaryons can be monitored over time by fluorescence microscopy combining staining with DAPI, antihuman LaminA/C, and F-actin/phalloidin (Fig. 4B.1.1C). For additional information, refer to Pereira et al. (2008).

Sort heterokaryons 32. At days 1, 2, and/or 3 aspirate medium, wash three times, each time with 10 ml CMF-PBS to remove cell debris and unfused lymphocytes. 33. Add 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes. Incubate 5 min at 37◦ C, 5% CO2 . 34. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate the trypsin. Dissociate cell clumps to single-cells by pipetting up and down. 35. Collect cells in 50-ml conical tubes. Centrifuge 5 min at 200 × g, room temperature. 36. Wash cells with 20 ml CMF-PBS and resuspend the cells in 5 ml of FACS buffer. 37. Pass the cells through a 70-μm cell strainer to remove clumps of cells. Transfer the cells to 5-ml polystyrene round-bottom tubes and put them on ice. 38. Sort the double-labeled population (heterokaryons, Fig. 4B.1.1B) using a FACS DiVa cell sorter (or equivalent) at 4◦ C.

Manipulation of Potency

4B.1.7 Current Protocols in Stem Cell Biology

Supplement 9

39. Wash recovered cells in 10 ml ice-cold CMF-PBS by centrifuging 5 min at 200 × g, 4◦ C. 40. Aspirate the supernatant completely and snap freeze the pellet in liquid N2 (day 1, 2, and 3) and store at −80◦ C.

Analyze expression in heterokaryons 41. After collection of the last sample (day 3), extract RNA from frozen samples (day 0, 1, 2, and 3) using RNA-BEE. Refer to Kingston et al. (1996) for a detailed procedure of RNA extraction. 42. Resuspend RNA pellets in 20 μl of DEPC-treated water and eliminate residual contaminating DNA by treatment with TURBO DNase-free kit. 43. Measure the RNA concentration with a Nanodrop spectrophotometer. 44. Reverse transcribe 1 to 3 μg of RNA using the Superscript first-strand synthesis system: dilute RNA in RNase-free water to a final volume of 11 μl and add 1 μl of 10 mM dNTP mix and 1 μl of Oligo (dT)12-18 . 45. Incubate 5 min at 65◦ C and on ice for 1 min. Add 1 μl of 0.1 M DTT, 4 μl of 5× first-strand buffer, 1 μl of RNaseOUT and 1 μl of 200 U/μl Superscript III. 46. Incubate samples 15 min at 25◦ C, 1 hr at 50◦ C,and 15 min at 75◦ C on a Dyad DNA engine. 47. Dilute cDNA 1/10 in water. Samples can be stored at −20◦ C for years.

48. Determine the abundance of human gene-specific transcripts in the heterokaryons at day 0, 1, 2, and 3 using quantitative RT-PCR (for review on real-time quantitative PCR, see Giulietti et al., 2001). We preferentially use SYBR Green Master Mix in a 35-μl reaction [2 μl of cDNA, 1.05 μl of combined primer pairs (10 μM each, Table 4B.1.1), 17.5 μl of Master Mix and 14.45 μl of water], using 96-well plates on a Chromo4 DNA engine under the following cycling conditions: 1 cycle: 40 cycles:

15 min 15 sec 30 sec 30 sec

95◦ C 94◦ C 60◦ C 72◦ C

(initial denaturation) (denaturation) (annealing) (extension).

Follow the PCR reaction with a plate reading. Perform each PCR reaction in triplicate. If designing new primer sets, ensure that the selected primers are species-specific and will therefore not cross-anneal with mouse sequences (Table 4B.1.1). Adjust cDNA dilution if the abundance of hGapdh is very different between samples.

49. Acquire data using the MJ Opticon Monitor 3 software. Use the comparative threshold method: for each primer pair set the threshold at the onset of the log-linear phase. The data can be exported to Excel spreadsheets. Calculate the amount of target genes normalized to the endogenous housekeeping gene (hGapdh) using the following equation: 2−C(T) × 1000 HeterokaryonBased Reprogramming for Pluripotency

where C(T) represents the threshold cycle at which fluorescence due to PCR products becomes detectable above background and C(T) is the C(T) of the target gene subtracted by the C(T) of the housekeeping gene (Fig. 4B.1.2).

4B.1.8 Supplement 9

Current Protocols in Stem Cell Biology

Current Protocols in Stem Cell Biology

Relative expression (/hGapdh)

0

0.01

0.02

0.03

0.04

0.05

4 3.5 3 2.5 2 1.5 1 0.5 0

16 14 12 10 8 6 4 2 0

d0

d1

d2

hTert

hDnmt3b

ES Oct4

ES WT

hOct4

d3

d0

d3

0.4

0.35 0.3 0.25 0.2 0.15 0.1 0.05 0

0.035 0.03 0.025 0.02 0.015 0.01 0.005 0

d1

d2

hRex1

hSox2

0.8

0.4

d0

d1

hHprt

hTle1

d2

hCripto

Time after fusion (days)

0

5

10

15

20

25

30

0

0.5

1

1.5

2

2.5

0

1.2

0.8

0

1.6

2

1.2

hNanog

1.6

2

d3

0

4

8

12

16

20

6 5 4 3 2 1 0

3.5 3 2.5 2 1.5 1 0.5 0

d0

d1

d2

hCD37

hCD45

hCD19

d3

Figure 4B.1.2 Kinetics of human lymphocyte reprogramming by mouse ES cells. Mouse ES cells expressing Oct4 (black bars), or in which Oct4 expression has been ablated (white bars, negative control; Niwa et al., 2000) are fused to hB lymphocytes and heterokaryons collected over the period of 3 days after cell fusion. The activation of human ES-specific genes (hOct4, hNanog, hCripto, hDnmt3b, hSox2, hTle1, hTert, and hRex1) and silencing of lymphocyte-specific genes (hCD19, hCD45, and hCD37) is quantified by qRT-PCR. Data is normalized to hGapdh and the expression of hHprt is added as a control gene. Error bars indicate the standard deviation (s.d.) of 2 to 3 independent experiments. Adapted from Pereira et al. (2008).

Manipulation of Potency

4B.1.9

Supplement 9

ALTERNATE PROTOCOL

GENERATION AND ANALYSIS OF INTER-SPECIES HETEROKARYONS WITHOUT FACS SORTING This protocol is used for the formation of heterokaryons between human somatic cells and mouse ES cells. This simplification of the Basic Protocol allows the selection of heterokaryons in culture based on a combination of drugs, overcoming the requirements for prelabeling and FACS sorting. Using drugs or specific cell lines carrying selective markers, it is possible to generate a population of heterokaryons suitable for gene expression analysis. Here, we give an example—the use of Ara-C in combination with ouabain for heterokaryon selection. This protocol is adequate when large numbers of cells are required for a specific analysis, for systematic cell fusion experiments, or when FACS sorting facilities are not easily accessible. Similar results have been obtained when this protocol was performed in parallel with the Basic Protocol.

Additional Materials (also see Basic Protocol) Ara-C (cytosine β-D-arabinofuranoside, Sigma, cat. no. C-1768) Ouabain (G-Strophanthin; Sigma, cat. no. O-3125) HAT (20 μM hypoxanthine, 0.08 μM aminopterine and 3.2 μM thymidine) media supplement (Sigma, cat. no. H0262-10VL), optional Puromycin (Sigma, cat. no. P9620), optional Generate inter-species heterokaryons without FACS sorting 1. Collect lymphocytes and ES cells following steps 1 to 6 from the Basic Protocol. Using this simplified protocol, cell labeling will not be necessary.

2. Mix cells and proceed with cell fusion. Follow steps 11 to 28 from the Basic Protocol. 3. Plate cells in 90-mm gelatin-coated dishes at ∼0.5 × 106 cells/cm2 with 15 ml ES/heterokaryon medium plus 1000 U/ml of LIF. Swirl the dish to ensure distribution over the entire plate surface. 4. Incubate 6 hr at 37◦ C, 5% CO2 . 5. Add Ara-C (10−5 M; 1 μl of 0.1 M stock per 10 ml of medium) and ouabain (10−5 M; 5.8 μl of 10 mg ml−1 stock per 10 ml of medium) to the dishes. This combination of drugs allows the selection of heterokaryons in culture. Proliferating ES cells are eliminated by Ara-C, a cytosine analog that inhibits DNA synthesis and therefore kills proliferating cells (nondividing cells, i.e., heterokaryons, will not be affected). Ouabain specifically kills human but not mouse cells (or inter-species heterokaryons). As an alternative to Ara-C, Hprt-/- ES cells (Hooper et al., 1987), which die in HAT selective medium, or puromycin-resistant human lymphocytes may be used.

6. Incubate 16 hr at 37◦ C, 5% CO2 . 7. Aspirate medium and wash once with 10 ml CMF-PBS to remove Ara-C. Add 10 ml of ES/heterokaryon medium plus 1000 U/ml LIF and ouabain (10−5 M). Change medium every day. 8. On day 1, 2, and 3, aspirate medium, wash three times, each time with 10 ml CMF-PBS to remove cell debris and unfused lymphocytes. 9. Add 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes. Incubate 5 min at 37◦ C, 5% CO2 . HeterokaryonBased Reprogramming for Pluripotency

10. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate trypsin. Collect cells in 15-ml conical tubes. Centrifuge 5 min at 200 × g, room temperature. 11. Wash recovered cells in 15 ml CMF-PBS by centrifuging 5 min at 200 × g, room temperature.

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12. Aspirate the supernatant completely and snap freeze the pellet in liquid N2 (day 1, 2, and 3) and store at −80 ◦ C. 13. Follow Basic Protocol steps 41 to 49 for RNA extraction, cDNA synthesis, and quantitative RT-PCR analysis.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

FACS buffer Add 1.5 ml of fetal bovine serum (FBS) to 48.5 ml calcium- and magnesium-free phosphate-buffered saline (CMF-PBS). Filter the solution using a 0.22-μm filter (Millipore, cat. no. SLGP033RS) and store up to 2 weeks at 4◦ C.

Gelatin, 0.1% (w/v) Incubate 2% (v/v) gelatin (Sigma, cat. no. G1393) for 15 min in a 37◦ C water bath to liquefy the solution. Add 25 ml of 2% (v/v) gelatin to 475 ml of calcium- and magnesium-free phosphate-buffered saline (CMF-PBS). Filter the solution with a bottle-top filter (0.22-μm; Millipore, cat. no. SCGPU05RE) and store up to 4 weeks at room temperature.

Gelatin-coated culture dishes Add enough 0.1% gelatin to cover the bottom of 90-mm dishes. Incubate for at least 20 min at 37◦ C. Before using, aspirate excess gelatin solution. We recommend coating culture dishes right before use.

hB cell medium RPMI-1640 (Invitrogen, cat. no. 31870-025) containing: 10% (v/v) heat-inactivated FBS 2 mM L-glutamine (Invitrogen, cat. no. 25030-123) 50 U/ml−1 penicillin and 50 μg/ml−1 streptomycin (Invitrogen, cat. no. 15140-122) Filter the medium with a 0.22-μm bottle-top filter Store up to 4 weeks at 4◦ C Mouse ES/heterokaryon medium Knockout DMEM (Invitrogen, cat. no. 10829-018) containing: 10% (v/v) fetal bovine serum (FBS) batch tested for ES cell culture 2 mM L-glutamine (Invitrogen, cat. no. 25030-123) 5 ml of non-essential amino acids (Invitrogen) 50 μM 2-mercaptoethanol (Invitrogen, cat. no. 31350-010) 50 U/ml−1 penicillin and 50 μg/ml−1 streptomycin (Invitrogen, cat. no. 15140122) Filter the medium with a 0.22-μm bottle-top filter Store up to 1 week at 4◦ C COMMENTARY Background information Reprogramming somatic cells to become ES-like is an important goal in cell replacement therapy since it affords the opportunity

to generate and use patient-specific ES-derived cells as grafts. Using this strategy, it would be possible to circumvent the problems of immune rejection that are likely to occur unless

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the recipient and donor stem cells are very closely matched. Several experimental reprogramming strategies to convert somatic cells to pluripotency have been outlined including nuclear transfer, cell fusion, and the forced expression of factors (Hochedlinger and Jaenisch, 2006; Yamanaka, 2007). These approaches have been instrumental for dissecting the re-establishment of pluripotency in a somatic cell at the cellular and molecular level. Retroviral-mediated expression of four transcriptional regulators—Oct4, Sox2, c-Myc, and Klf4—was shown to drive mouse and human fibroblasts into an ES-like state, albeit at low frequency (Jaenisch and Young, 2008; Yamanaka, 2007). These studies have illustrated the importance of several factors for reprogramming, but they also suggested that additional ones might be needed for efficient conversion to pluripotency. Reprogramming can also be achieved by cellular fusion, a process that occurs spontaneously in vitro (Ying et al., 2002), in vivo (Weimann et al., 2003), and experimentally using specific agents (Terranova et al., 2006; Pereira et al., 2008). For example, fusion of differentiated cells with pluripotent ES cells induces the expression of pluripotency-associated markers in the hybrid cells and chromatin remodeling at specific sites in the somatic cell genome (Tada et al., 1997; Cowan et al., 2005). To study reprogramming in heterokaryons and hybrid cells it is important to be able to distinguish and verify events that occur in the donor nucleus from those of the recipient. Inter-species heterokaryons formed between human lymphocytes and mouse ES cells therefore provide a tractable approach to study direct reprogramming to pluripotency.

Critical Parameters and Troubleshooting Troubleshooting advice can be found in Table 4B.1.2.

Anticipated Results

HeterokaryonBased Reprogramming for Pluripotency

A total of 1 × 108 mouse ES cells and 1 × 108 human B lymphocytes are adequate to sort at least 300,000 heterokaryons per time point (day 1, 2, and 3). This allows the parallel gene expression analysis by qRT-PCR of

human pluripotency-associated genes (hOct4, hNanog, hCripto, hDnmt3b, hSox2, hTle1, hTert, and hRex1), lymphocyte-specific genes (hCD19, hCD45, and hCD37), and housekeeping controls (hGapdh, hHprt). As shown in Figure 4B.1.2, fusion of human lymphocytes with wild-type ES cells (black bars) results in the increased expression of human pluripotency-associated genes over this 3-day period. Pluripotency-associated gene activation is mirrored by a reduction in expression of several human lymphocyte-associated genes after heterokaryon formation. As a control, ES cells lacking Oct4 expression [essential transcription factor for ES cell self-renewal and reprogramming ability (Niwa et al., 2000; Pereira et al., 2008)] are fused to human B lymphocytes (white bars) and the activation of human pluripotency-associated genes is impaired. Reprogramming kinetics should be representative of at least two independent experiments.

Time Considerations Expansion of human B lymphocytes and mouse ES cells (steps 1 and 2) takes 7 days. Usually five 90-mm dishes of confluent ES cells and one 175-cm2 flask of lymphocytes are enough to get 1 × 108 cells each cell-type. On day 0, collection of cell types, counting, and cell labeling (steps 3 to 10) will take between 1 and 1.5 hr. The cell fusion procedure and heterokaryon culture (also performed on day 0; steps 11 to 31) will take ∼30 to 45 min. Pause point: cells are left overnight in culture. Days 1 through 3, including reprogramming kinetics and collection of samples (steps 32 to 40) will require 3 days. Cell sorting duration may vary depending on the instrument and cell number, usually 2 to 3 hr per sample. Pause point: cell pellets can be stored at −80◦ C for months. On day 4, RNA extraction, DNase treatment, and reverse transcription (steps 41 to 47) will take 5 hr. Pause point: reverse transcription can be set up in PCR strip tubes and left overnight including a final step at 4◦ C. On day 5, real-time PCR set up, run, and analysis (steps 48 and 49) will require 5 hr. Setting up a real time PCR plate will take ∼45 min and the PCR run takes 3 to 4 hr, which can be left running overnight.

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Table 4B.1.2 Troubleshooting Guide to Heterokaryon Formation and Analysis

Problem

Possible cause

Solution

Heterokaryons are not forming

Poor quality of fused cells

Make sure that both ES cells and B lymphocytes are well maintained in culture. Change medium the day before cell fusion.

PEG is being diluted

Dilution of PEG will result in decreased fusion efficiency. Before addition of PEG, remove PBS completely including drops on the surface of tube.

Cell fusion temperature is not optimal

Always warm the cell pellet and PEG in a 37◦ C water bath. Check the water bath temperature. If necessary, add PEG dropwise while keeping the tube in the water bath.

Mechanical dissociation

A period of cell contact is required to complete cell fusion. Handle heterokaryons carefully. Do not mix or pipet heterokaryons vigorously.

Cell labeling is not optimal

Homogeneous labeling with minimum toxicity has to be optimized. Reduce both dye concentration and labeling time.

PEG treatment

Upon PEG treatment a balance between cell death and fusion efficiency has to be found. Reduce time in PEG or predilute PEG in PBS to decrease cell death.

Osmotic shock after PEG treatment

After PEG treatment add DMEM dropwise slowly over a longer period of time (4 ml over 5 min). Make sure to gently rock the tube continuously to slowly dilute PEG.

Cell sorting conditions are not optimal

Check the purity of sorted population in a control experiment by immnunofluorescence. Exclude doublets of cells when sorting heterokaryons.

Low number of heterokaryons sorted

Increase the number of starting cells for cell fusion or optimize cell ratio to increase efficiency.

Poor RNA quality

Check quality of RNA on a gel. If RNA is degraded, skip DNase treatment. Refer to Kingston et al. (1996) for troubleshooting.

Poor quality of ES cell line

Test another mouse ES cell line, or clone, for cell for heterokaryon-based reprogramming.

cDNA is too diluted

The activation of genes will occur gradually and will be detected at late cycles. Use concentrated cDNA to start with and dilute according hGapdh levels.

Refractory human somatic cell line

Some cell types can be more or less prone to reprogramming. Use another cell type or cell line. We have successfully reprogrammed several human EBV–transformed B cell clones.

Sample contamination with unfused hB lymphocytes

Add ouabain to the heterokaryon medium after cell fusion. Wash dishes very well with PBS to remove lymphocytes. Decrease size of gate when sorting cells to avoid the collection of DiI+ DiD− cells.

Excessive cell death

hGapdh levels very low in heterokaryons

Pluripotency-associated genes are not detected

Lymphocyte-associated genes are not silenced

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Literature Cited Cowan, C.A., Atienza, J., Melton, D.A., and Eggan, K. 2005. Nuclear reprogramming of somatic cells after fusion with human embryonic stem cells. Science 309:1369-1373. Giulietti, A., Overbergh, L., Valckx, D., Decallonne, B., Bouillon, R., and Mathieu, C. 2001. An overview of real-time quantitative PCR: Applications to quantify cytokine gene expression. Methods 25:386-401. Hochedlinger, K. and Jaenisch, R. 2006. Nuclear reprogramming and pluripotency. Nature 441:1061-1067. Hooper, M., Hardy, K., Handyside, A., Hunter, S., and Monk, M. 1987. HPRT-deficient (LeschNyhan) mouse embryos derived from germline colonization by cultured cells. Nature 326:292295. Jaenisch, R. and Young, R. 2008. Stem cells, the molecular circuitry of pluripotency and nuclear reprogramming. Cell 132:567-582. Kingston, R.E., Chomczynski, P., and Sacchi, N. 1996. Guanidine methods for total RNA preparation. Curr. Protoc. Mol. Biol. 36:4.2.1-4.2.9. Niwa, H., Miyazaki, J., and Smith, A.G. 2000. Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat. Genet. 24:372-376. Pereira, C.F., Terranova, R., Ryan, N.K., Santos, J., Morris, K.J., Cui, W., Merkenschlager, M., and Fisher, A.G. 2008. Heterokaryon-based reprogramming of human B lymphocytes for pluripotency requires Oct4 but not Sox2. PLoS Genet. 4:e1000170.

Silva, J., Chambers, I., Pollard, S., and Smith, A. 2006. Nanog promotes transfer of pluripotency after cell fusion. Nature 441:997-1001. Stadtfeld, M., Maherali, N., Breault, D.T., and Hochedlinger, K. 2008. Defining molecular cornerstones during fibroblast to iPS cell reprogramming in mouse. Cell Stem Cell 2:230240. Tada, M., Tada, T., Lefebvre, L., Barton, S.C., and Surani, M.A. 1997. Embryonic germ cells induce epigenetic reprogramming of somatic nucleus in hybrid cells. EMBO J. 16:65106520. Tada, M., Takahama, Y., Abe, K., Nakatsuji, N., and Tada, T. 2001. Nuclear reprogramming of somatic cells by in vitro hybridization with ES cells. Curr. Biol. 11:1553-1558. Terranova, R., Pereira, C.F., Du Roure, C., Merkenschlager, M., and Fisher, A.G. 2006. Acquisition and extinction of gene expression programs are separable events in heterokaryon reprogramming. J. Cell Sci. 119:2065-2072. Weimann, J.M., Johansson, C.B., Trejo, A., and Blau, H.M. 2003. Stable reprogrammed heterokaryons form spontaneously in Purkinje neurons after bone marrow transplant. Nat. Cell Biol. 5:959-966. Yamanaka, S. 2007. Strategies and new developments in the generation of patient-specific pluripotent stem cells. Cell Stem Cell 1:39-49. Ying, Q.L., Nichols, J., Evans, E.P., and Smith, A.G. 2002. Changing potency by spontaneous fusion. Nature 416:545-548.

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Imaging Neural Stem Cell Fate in Mouse Model of Glioma

UNIT 5A.1

Khalid Shah1 1

Massachusetts General Hospital, Harvard Medical School, Charlestown, Massachusetts

ABSTRACT This unit describes a protocol for following the fate of stem cells in real time in a mouse model of glioma. Stem cells and tumor cells can be transduced with lentiviral vectors bearing two different luciferases, firefly luciferase (Fluc) and Renilla (Rluc) luciferase, respectively. With the cells labeled in this manner, bioluminescence imaging can be used to study the fate of stem cells in glioma-bearing brains in vivo. Curr. Protoc. Stem Cell C 2009 by John Wiley & Sons, Inc. Biol. 8:5A.1.1-5A.1.11.  Keywords: neural stem cell r bi-modal vector r luciferase r fluorescent proteins r glioma r in vivo imaging

INTRODUCTION Several studies have demonstrated the effectiveness of neural stem cell (NSC) transplantation in the treatment of neurodegenerative diseases, including spinal cord injury and brain tumors (Snyder and Macklis, 1995; Ehtesham et al., 2002; Lindvall et al., 2004; Hofstetter et al., 2005; Iwanami et al., 2005; Shah et al., 2005). This unit describes a protocol for simultaneously imaging the fate of engineered NSC and glioma cells in a mouse glioma model. NSC and glioma cells transduced with lentiviral vectors bearing different combinations of fluorescent and bioluminescent proteins can be grown as monolayers and maintained over several passages. The unit begins with a method for transducing NSC and glioma cells with bimodal lentiviral vectors for stable expression of these fluorescent and bioluminescent markers in vitro, followed by transplantation of fluorescent and bioluminescent glioma cells and NSC in mice, and, finally, sequential bioluminescent imaging of NSC fate and glioma progression in mice. The integration of different combinations of bioluminescent and fluorescent proteins into NSC and glioma cells makes it possible to distinguish different populations of cells after intracranial transplantation. Lentiviral vector transduction of cells is followed by cell sorting, which is necessary to obtain a pure population of different fluorescent cell types. The protocol details viral transduction, surgical preparation, craniotomy, cell implantation, animal recovery, and imaging procedures to study stem cell kinetics and migration to malignant brain tumors. NOTE: All solutions and equipment coming into contact with live cells must be sterile. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5%, CO2 incubator unless otherwise specified. NOTE: Viral transductions on human stem cells and glioma cells and cell culture procedures are performed in a biosafety level (BL)-2 facility in a laminar-flow hood.

ENGINEERING STEM CELL AND GLIOMA LINES This protocol is used for transducing human NSC and human glioma cells with lentiviral vectors bearing bioluminescent and fluorescent markers for stable expression of these markers in vitro and in vivo. Both cell types are transduced with lentiviral vectors bearing unique combinations of fluorescent and bioluminescent markers, and cells are sorted by cell sorter. Current Protocols in Stem Cell Biology 5A.1.1-5A.1.11 Published online March 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a01s8 C 2009 John Wiley & Sons, Inc. Copyright 

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Materials Human neural stem cells (NSC; Rubio et al., 2000) NSC culture medium (see recipe) 0.25% (w/v) trypsin/EDTA (Invitrogen) Human glioma cells (Gli36; Shah et al., 2004) Glioma cell culture medium: DMEM containing 10% FBS and 1× penicillin/streptomycin Plasmid for NSC cells: lentiviral plasmid bearing a fusion between GFP and Fluc (GFP-Fluc; Shah et al., 2008) Plasmid for glioma cells: lentiviral plasmid bearing a fusion between Rluc and DsRed2 (Rluc-DsRed2; Shah et al., 2008) Phosphate-buffered saline (PBS; e.g., Invitrogen) NSC culture medium (see recipe) containing 8 μg/ml polybrene (add from 8 mg/ml polybrene stock in PBS; Fisher) Glioma cell culture medium (see above) containing 8 μg/ml polybrene (add from 8 mg/ml polybrene stock in PBS; Fisher) 5-cm culture dishes (Corning) Fluorescence microscope with appropriate filters for GFP and rhodamine Cell sorter (e.g., FACScalibur from BD Biosciences) Additional reagents and equipment for fluorescence-activated cell sorting (Robinson et al., 2009) Culture cells and lines 1a. For human fetal neural stem cell line: Culture human fetal neural stem cell line (NSC), derived from the human diencephalic and telencephalic regions of 10 to 10.5 weeks gestational age from an aborted human Caucasian embryo, in NSC culture medium in a 5-cm culture dish at 37◦ C in a humidified incubator, to 70% to 80% confluency. These cells are grown as monolayers and are passaged every 4 days by trypsinizing cells in 0.25% trypsin/EDTA. Cells are centrifuged 10 min at 300 × g and plated at 20% density in NSC culture medium. The in vitro and in vivo properties of NSC (including the absence of transformation, clonality, multipotency, stability, and survival) have been described in detail elsewhere (Rubio et al., 2000; Villa et al., 2004; Navarro-Galve et al., 2005).

1b. For human glioma cell line: Culture Gli36, a human glioma cell line whose in vitro and in vivo characteristics have been described elsewhere (Shah et al., 2004, 2005) in glioma cell culture medium at 37◦ C to 70% to 80% confluency. Glioma cells grow as monolayers and are passaged every 4 days by trypsinizing cells in 0.25% trypsin/EDTA. Cells are seeded at 20% density in glioma cell culture medium.

2. When cells reach 70% to 80% confluency, subculture cells at a 1:4 (NSC) or 1:5 (glioma cell) ratio.

Prepare lentiviral vectors 3. Use the CS-CGW transfer plasmid–based lentiviral vector system (Miyoshi et al., 1998) to create lentiviral transfer vectors bearing fusions between Renilla luciferase (Rluc) and Discosoma Red (DsRed2) proteins (LV-Rluc-DsRed2) and lentiviral vectors bearing fusions between firefly luciferase (Fluc) and green fluorescent protein (GFP; LV-GFP-Fluc). Imaging Neural Stem Cell Fate in Mouse Model of Glioma

The construction of LV-GFP-Fluc and LV-Rluc-DsRed2 is described in detail elsewhere (Shah et al., 2008).

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4. Amplify the cDNA sequences encoding GFP-Fluc fusion and Rluc-DsRed2 fusion by PCR (Kramer and Coen, 2001) and ligate in-frame into NheI/XhoI-digested CS-CGW plasmid (Shah et al., 2008). 5. Produce lentiviral vectors (Shah et al., 2008). 6. Titer the viruses (Shah et al., 2008) and store in PBS at −80◦ C.

Perform viral transduction and cell sorting For NSC 7a. Plate NSC at 60% confluency in a 5-cm dish. 8a. At a time point 18 hrs later, transduce NSC with LV-GFP-Fluc (at MOI = 1) in NSC culture medium containing 8 μg/ml polybrene. 9a. Confirm viral transduction by visualizing cells for GFP expression by fluorescence microscopy 36 to 48 hr after transduction. 10a. At 72 hr after transduction, perform single-cell sorting based on GFP fluorescence, using a cell sorter (also see Robinson et al., 2009) to obtain a monoclonal cell populations. Culture sorted cells in NSC culture medium. We use BD FACScalibur cell sorter (BD Biosciences).

For glioma cells 7b. Plate glioma cells at 40% confluency in a 5-cm dish. 8b. 18 hr later, transduce glioma cells with LV-Rluc-DsRed2 (at MOI = 1) in glioma cell culture medium containing 8 μg/ml polybrene. 9b. Confirm viral transduction by visualizing cells for DsRed2 expression by fluorescence microscopy 48 hr after transduction. 10b. At 72 hr after transduction, trypsinize cells using trypsin/EDTA, wash the cells twice with PBS (each time centrifuging 10 min at 300 × g), and perform single-cell sorting based on rhodamine fluorescence using a cell sorter (also see Robinson et al., 2009) to obtain monoclonal cell populations. Culture sorted cells.

BIOLUMINESCENCE IMAGING IN CULTURE This protocol is used for bioluminescence imaging of NSC and glioma cells expressing different combinations of bioluminescent and fluorescent markers in vitro (see Fig. 5A.1.1).

SUPPORT PROTOCOL

Materials NSC and glioma cells bearing bioluminescent and fluorescent markers (Basic Protocol 1) NSC culture medium (see recipe) 150 mg/ml D-luciferin stock (firefly luciferase substrate; Biotium, cat. no. 10110-1; http://www.biotium.com) in PBS Glioma cell culture medium: DMEM containing 10% FBS 1 mg/ml coelenterazine stock (substrate for Renilla luciferases; Biotium, cat. no. 10102-2; http://www.biotium.com) in ethanol 48- or 96-well clear-bottom black-walled plate Bioluminescence imaging system with IVIS-200 or IVIS-100 (Caliper; http://www.caliperls.com/) or similar bioluminescence imaging system Genetic Manipulation of Stem Cells

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Figure 5A.1.1 Fluorescence and bioluminescence characteristics of NSC (A) and glioma cells (B) in culture. NSC and glioma cells were transduced in culture with LV-GFP-Fluc and Lv-RlucDsRed2, respectively, at MOI = 1, and visualized for GFP (A) or DsRed2 (B) ßuorescence. (C,D) Different concentrations of NSC expressing GFP-Fluc (1.0–1.5 × 105 ) and glioma cells (1.5–6 × 105 ) expressing Rluc-DsRed2 were plated, and, 12 hr later, cells were incubated with 150 μg/ml D-luciferin or 1 μg/ml of coelenterazine and imaged under the CCD with a scan time of 1 min. MagniÞcation, 20×. Adapted from Shah et al. (2008), with permission from Society for Neuroscience.

To image the bioluminescence of transduced NSC 1a. Using a black-walled, clear-bottom 96-well tissue culture plate, seed NSC at several densities spanning 1000 to 10,000 cells per well in 100 μl NSC culture medium, to determine the correlation between the number of transduced NSC and the firefly luciferase bioluminescence signal. Incubate. 2a. At a time point 18 to 24 hr later add D-luciferin (substrate for firefly luciferase) to the culture medium at a 1/10 volume, for a final concentration of 0.15 mg/ml, using a multichannel pipettor. Dilute from a 150 mg/ml D-luciferin stock.

3a. Rock the plate and take images in bioluminescence imager with the appropriate exposure. For firefly luciferase, peak light production from intact cells occurs ∼10 min after substrate addition.

To image bioluminescence imaging of transduced glioma cells 1b. Using a black-walled, clear-bottom 96-well tissue culture plate, seed glioma cells at several densities ranging from 1000 to 10,000 cells per well in 100 μl glioma cell culture medium to determine the correlation between number of transduced glioma cells and Renilla luciferase bioluminescence signal. Incubate. Imaging Neural Stem Cell Fate in Mouse Model of Glioma

2b. At a time point 18 to 24 hr later, add coelenterazine (substrate for Renilla luciferase) to the culture medium at a 1/10 volume, for a final concentration of 0.1 μg/ml, using a multichannel pipettor.

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The 1 mg/ml ethanol stock of coelenterazine is first diluted to an appropriate concentration with PBS before being added to the culture medium.

3b. Rock the plate and take images in bioluminescence imager with the appropriate exposure. Peak bioluminescence from Renilla luciferases occurs rapidly within the first minute after adding coelenterazine.

CELL TRANSPLANTATION AND IMAGING This protocol is used for transplantation and subsequent imaging of NSC and glioma cells expressing different combinations of bioluminescent and fluorescent markers in mice. It also describes the dual imaging of NSC fate and glioma progression in the mouse glioma model.

BASIC PROTOCOL 2

NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Committee for Ethical Animal Care and Use (IACUC) and must conform to government regulations for the care and use of laboratory animals. NOTE: Mouse surgical procedures are performed in a surgical room designated for animal surgeries. Proper aseptic techniques should be used accordingly.

Materials SCID mice (6-to 8-weeks-old; Charles River Laboratories) Anesthetics: ketamine and xylazine (also see Donovan and Brown, 1998) Betadine solution (Bruce Medical; http://www.brucemedical.com/) 70% isopropyl alcohol (Fisher) Phosphate-buffered saline (PBS), sterile Gli36-Rluc-DsRed2 glioma cells (Basic Protocol 1) Bone wax (Ethicon) Coelenterazine (100 μg/animal in 150 μl saline; Biotium, cat. no. 10102-2) D-luciferin (150 μg/g body weight in 150 μl saline; Biotium, cat. no. 10110-1) Animal shaver Stereotaxic frame (Harvard Apparatus, cat. no. 726049) Stereo dissecting microscope: variable magnification (1 to 4.5; Nikon) Fine scissors (Fine Science Tools, cat. no. 14084-08) Forceps, angled and straight and ultrafine angled (Fine Science Tools) Cotton-tipped applicators Hand-held micro-drill (Fine Science Tools, cat. no. 18000-17) with 0.45-mm round drill burr (VWR) 10-μl Hamilton gastight 1701 syringe with 26-G needle 4–0 vicryl sutures or surgical staples Bioluminescence imaging system with IVIS-200 or IVIS-100 (Caliper; http://www.caliperls.com/) or similar bioluminescence imaging system Additional reagents and equipment for anesthesia of mice (Donovan and Brown, 1998) Anesthetize the animal 1. Grasp the animal firmly with one hand and anesthetize by injecting ketamine and xylazine intraperitoneally (Donovan and Brown, 1998). The ideal dosage for each animal will vary primarily based upon the animal’s body mass (120 mg/kg ketamine and 16 mg/kg xylazine) and age.

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Figure 5A.1.2

Anesthetized mouse in a stereotaxic device being implanted with glioma cells.

2. Use the toe-pinch method to assess the level of sedation by firmly applying pressure to the animal’s toe pads and observing whether or not there is a demonstration of a pain response by the animal. Also monitor breathing and posture. 3. Secure animal on a stereotactic head frame placed under a stereo dissecting microscope and shave dorsal surface of the animal’s head (see Fig. 5A.1.2). 4. Disinfect the shaved area by applying two alternating coatings with Betadine and 70% isopropyl alcohol. 5. Using scissors and forceps, remove the skin from the disinfected region and use a dry cotton swab to completely remove the periosteum membrane from the exposed skull surface. Keep the skull moist by frequent application of sterile PBS following the removal of the periosteum. 6. For glioma cell implantations, use a handheld micro-drill to drill through the bone at the location of the proposed implantation site until the cortical surface is exposed.

Implant tumor cells 7. Place 4 to 5 μl of Gli36-Rluc-DsRed2 glioma cells (100,000 cells) in a 10-μl 26-G Hamilton Gastight 1701 syringe and insert the needle to a specified depth into the left frontal lobe. In our experiments we have used the following stereotactic coordinates: 2.5 mm lateral and 0.5 mm caudal to bregma; depth 2.5 mm from dura.

8. Implant cells over a period of 4 min with 30-sec intervals. Care should be taken to consistently implant tumors at the same location and depth to facilitate bioluminescence interpretation from within this relative point source.

9. After implantation is complete, wait for 5 min and remove needle over a period of 10 min with intervals of 1 min. 10. Seal the burrow hole with bone wax and close the wound with 4–0 vicryl sutures or surgical staples. Imaging Neural Stem Cell Fate in Mouse Model of Glioma

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Figure 5A.1.3 NSC migrate into gliomas in vivo. Bioluminescence imaging of mice implanted with GFP-Fluc expressing NSC in mice with established Rluc-DsRed2 gliomas. Fluc images of mice on day 3 (A), day 7 (B), and day 10 (C), and Rluc image on day 10 (D). Adapted from Shah et al. (2008) with permission from Society for Neuroscience.

Image in vivo tumor cell bioluminescence Imaging can be performed 24 hr after cell implantation. 11. Anesthetize mouse by injecting the appropriate dose of ketamine and xylazine intraperitoneally (see step 1). 12. Use the toe-pinch method to assess the level of sedation by firmly applying pressure to the animal’s toe pads and observing whether or not there is a demonstration of a pain response by the animal. Also monitor breathing and posture. It is slightly more difficult, yet equally important, to monitor anesthesia during imaging as during surgery. During extended time-course sessions, imaging may be jeopardized by a possible toe-pinch reaction and it may be more appropriate to monitor the animal’s breathing and posture.

13. First, acquire a surface image of each animal using dim polychromatic illumination. Next, measure the spatial distribution of luciferase activity within the mouse brain by photon count recording using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3), according to the manufacturer’s instructions. 14. Image mice for Rluc activity by injecting 100 μg coelenterazine (in 150 μl saline) intravenously via the tail vein and record photon counts 5 min later over a 5-min period using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3) according to the manufacturer’s instructions.

Implant stem cells 15. Anesthetize the same animals implanted with glioma cells with the appropriate dose of ketamine and xylazine (see step 1). 16. Secure on a stereotactic head frame placed under a stereo dissecting microscope. 17. Using a handheld micro-drill, drill hole in the contralateral, right frontal lobe at the following coordinates: 2.5 mm lateral and 0.5 mm caudal to bregma; depth 2.5 mm from dura. Depending on the migrating ability and speed of migration, NSC can be placed at any distance from the gliomas in order to assess migration of NSC to gliomas in the brain. In our studies, we have placed the NSC in the contralateral right frontal lobe of the glioma-bearing mice in order to follow migration of NSC toward gliomas.

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18. Place 4 to 5 μl of NSC expressing GFP-Fluc (NSC-GFP-Fluc; 500,000 cells) in a 10-μl 26-G Hamilton Gastight 1701 syringe and implant cells over 4 min with 30-sec intervals. 19. Wait 5 min and withdraw the syringe over 10 min, with 1-min intervals.

Image in vivo stem cell bioluminescence 20. To image mice for firefly luciferase (Fluc) activity, inject the mice intraperitoneally with 150 μg/g body weight D-luciferin (in 150 μl saline). 21. Acquire images 10 min after D-luciferin administration over a period of 5 min. 22. Measure the spatial distribution of luciferase activity within the brain of the animal by recording photon counts using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3), according to the manufacturer’s instructions. Mice can be imaged every day for Fluc and Rluc activity. Typical exposure times vary between 1 and 10 min. If imaging for both, screen for Rluc activity and then Fluc activity. Allow a 24-hr period between imaging sessions to make sure there is no residual luciferase activity from the previous session.

Allow the animal to recover 23. Observe the animal for recovery. Make certain the animal is restrained and that it cannot cause harm to itself. When the animal is maintaining its own normal body temperature and has a reflexive response to toe-pinch stimulation, return it to a clean and unoccupied cage. For the most part, the animal should survive the procedure despite the absence of an external heat source. The usual recovery time for this procedure can range from 2 to 12 hr. If the animal has not resumed normal grooming and eating behavior beyond this time frame, it may require additional medical attention or euthanasia.

Analyze data 24. Use the software accompanying the imaging equipment to perform the region of interest (ROI) analysis. In our studies, following data acquisition, post-processing and visualization is performed using a home-written program with image display and analysis suite developed in IDL (Research Systems Inc.). Regions of interest are defined using an automatic intensity contour procedure to identify bioluminescence signals with intensities significantly greater than the background. The mean, standard deviation, and sum of the photon counts in these regions are then calculated. For visualization purposes, the bioluminescence images are fused with the corresponding white-light surface images as a transparent pseudocolor overlay, permitting correlation of areas of bioluminescence activity with anatomy. Maintaining a standard region of interest within an experiment (or series of experiments) is important to facilitate comparison of mouse imaging data.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

NSC culture medium

Imaging Neural Stem Cell Fate in Mouse Model of Glioma

DMEM/F-12 (Invitrogen) supplemented with: 0.6% (w/v) D-glucose (Sigma-Aldrich) 0.5% (w/v) AlbuMax (Life Technologies) 0.5% (w/v) L-glutamine (Life Technologies) 20 ng/ml recombinant human FGF (R & D Systems) continued

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20 ng/ml recombinant human EGF (R & D Systems) 1× N2 supplement (Invitrogen) 1% (w/v) nonessential amino acids (Cellgro) 1 mM sodium pyruvate (Cellgro) 26 mM sodium bicarbonate) COMMENTARY Background Information Neural stem cells (NSC) are defined by their ability to self-renew and give rise to mature progenitors of neural lineages. The ability of NSC to migrate to diseased areas of the brain has been documented (Snyder and Macklis, 1995: Aboody et al., 2000; Tang et al., 2003; Shah et al., 2005). Their capacity to differentiate into all neural and glial phenotypes (Gage, 2000) provides a powerful tool for targeting the treatment of both diffuse and localized neurologic disorders. Several studies have demonstrated the effectiveness of NSC transplantation in the treatment of neurodegenerative diseases, including spinal cord injury and brain tumors (Snyder and Macklis, 1995; Ehtesham et al., 2002; Lindvail et al., 2004; Hofstetter et al., 2005; Iwanami et al., 2005; Shah et al., 2005). Taking advantage of their homing properties, NSC have also been modified to deliver selective anti-neoplastic proteins (Ehtesham et al., 2002; Shah et al., 2005), although with mixed results. While these studies demonstrate the feasibility of NSC-based therapy, cellular delivery of therapeutic proteins via NSC grafts will likely require long-term transgene expression. In vivo assays, which permit rapid assessment of the fate of transplanted stem cells, will be useful in designing future stem-cell-based therapies. Bioluminescence imaging exploits the emission of visible photons at specific wavelengths based on energy-dependent reactions catalyzed by luciferases. It is a powerful method for detecting and quantifying the spatial and temporal occurrence of cellular and molecular events and can be efficiently used for longitudinal comparison of cell survival and migration. Luciferases from Renilla (Rluc) and firefly (Fluc) have different substrates, coelenterazine and D-luciferin, respectively, and can be imaged in tumors in the same living mouse with kinetics of light production being separable by timed injections of these two substrates (Shah et al., 2005). This dual-imaging approach has direct applications in studying gene expression from vectors and simultaneously monitoring therapeutic effects in vivo.

Critical Parameters and Troubleshooting An efficient and robust way to follow cells both in culture and in vivo is to transduce them with lentiviral vectors expressing fusions of bioluminescent and fluorescent marker genes. These vectors have the ability to integrate transgenes into the genome of dividing and nondividing cells (Naldini et al., 1996) and provide means of efficient long-term expression in cells and their progeny without using any antibiotic selection marker. The fluorescent marker serves to determine the efficiency of transduction, and, in conjunction with the bioluminescent marker, serves as an in vivo cell-tracking protein. Furthermore, the expression of fluorescent markers in different cell populations also aids in performing pathological analysis on tissue sections in sacrificed animals. Knowing the depth and optical properties of the tissue through which the light will pass is essential in calculating numbers of cells needed to obtain a detectable signal. Generally, firefly luciferase light will be attenuated approximately 10-fold for each centimeter of tissue, but optically dense tissues such as liver will attenuate light much more than skin, bone, or lung. Thus, the number of luciferaseexpressing cells and their localization within the body is critical in obtaining a detectable signal to follow fate of cells in vivo; the deeper the tumors are within the body, or the deeper the intracranial tumor, the greater the signal attenuation. For example, in subcutaneous tumors, cell numbers as low as 1000 firefly luciferase–expressing cells can be detected. Also, D-luciferin has more favorable biodistribution than coelenterazine, and an intraperitoneal injection of luciferin is much more reproducible than the tail vein injection that is required for delivering coelenterazine. Transplanting cells expressing Fluc and Rluc in various sites, using various gene delivery vectors and transgenic models, demonstrates the high accessibility of D-luciferin (Fluc substrate) to various tissues, including the brain. On the other hand, coelenterazine

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is also accessible to many tissues because of its diffusable nature (Lorenz et al., 1996), but its distribution in the intact brain is limited by drug-transport proteins, which can hinder in vivo imaging of Renilla luciferase (Pichler et al., 2004). This problem may be overcome by injecting mice with a blood-brain barrier (BBB) disrupter, e.g., mannitol. Also, mouse fur attenuates and scatters light, and this effect is most pronounced in black mice. This problem may be overcome by using nude mice or shaving animals over the region(s) of interest for imaging. Luciferase imaging in mice offers the possibility of imaging mice serially. To perform repetitive imaging of mice, the user should take into account that luciferase levels in mice peak ∼10 min after intraperitoneal. injection, then decline slowly to background levels by 6 to 8 hr post injection (Paroo et al., 2004). Coelenterazine has a more rapid kinetic course in mice. Therefore, maximum imaging signal for Renilla luciferases is obtained immediately after injecting coelenterazine through intravenous or intra-cardiac routes (Bhaumik and Gambhir, 2002). For imaging two different molecular events simultaneously, for example, stem cell fate and glioma volumes in the same mouse, it is advisable to image Renilla luciferase activity first, and then image firefly luciferase activity. Bioluminescence signal from mice implanted with NSC-expressing bioluminescent proteins in the brain varies with the presence and absence of tumors, and in different mice. We have previously shown that human neural stem cell survival is much improved in SCID mice as compared to nude mice (Shah et al., 2008), which could be attributed to the fact that SCID mice (lacking functional T and B cells) are more immune-compromised than nude mice (which lack functional T cells only), and this may implicate immune rejection as a factor in NSC survival in the brain. Our studies also reveal the persistence of NSC in the brains of tumor-bearing mice as compared to normal mice, implying that glioma cells or host response may modulate human stem cell survival either through secretion of growth factors or by inhibition of molecules involved in foreign cell rejection. While designing experiments for imaging human stem cell fate in mouse tumor models, the choice of mouse and the tumor cell type should be taken into consideration.

Anticipated Results The protocols in this unit generate useful information on the fate of NSC in a mouse model of glioma, and are suitable for a number of other disease models. Both stem cells and glioma cells can be easily transduced with lentiviral vectors, and bioluminescence imaging can be used to study the fate of stem cells in different disease models in vivo. Glioma cell survival is higher than NSC survival in mice. Furthermore, we have shown that the presence of glioma cells improves the survival of NSC in the brain.

Time Considerations It takes 1 week for glioma cells and 2 weeks for NSC to grow before they are transduced with lentiviral vectors. Glioma cells are implanted 2 to 3 days after lentiviral transduction, and transduced NSC are implanted 3 to 4 days after glioma cell implantation. Both glioma cells and NSC can be followed in real time in vivo for a period of 3 to 4 weeks before glioma growth results in the mortality of animals.

Literature Cited Aboody, K.S., Brown, A., Rainov, N.G., Bower, K.A., Liu, S., Yang, W., Small, J.E., Herrlinger, U., Ourednik, V., Black, P.M., Breakefield, X.O., and Snyder, E.Y. 2000. Neural stem cells display extensive tropism for pathology in adult brain: Evidence from intracranial gliomas. Proc. Natl. Acad. Sci. U.S.A. 97:12846-12851. Bhaumik, S. and Gambhir, S.S.. 2002. Optical imaging of Renilla luciferase reporter gene expression in living mice. Proc. Natl. Acad. Sci. U.S.A. 99:377-382. Donovan, J. and Brown, P. 1998. Anesthesia. Curr. Protoc. Immunol. 27:1.4.1-1.4.5. Ehtesham, M., Kabos, P., Gutierrez, M.A., Chung, N.H., Griffith, T.S., Black, K.L., and Yu, J.S. 2002. Induction of glioblastoma apoptosis using neural stem cell-mediated delivery of tumor necrosis factor-related apoptosis-inducing ligand. Cancer Res. 62:7170-7174. Gage, F.H. 2000. Mammalian neural stem cells. Science 287:1433-1438. Hofstetter, C.P., Holmstrom, N.A., Lilja, J.A., Schweinhardt, P., Hao, J., Spenger, C., Wiesenfeld-Hallin, Z., Kurpad, S.N., Frisen, J., and Olson, L. 2005. Allodynia limits the usefulness of intraspinal neural stem cell grafts: Directed differentiation improves outcome. Nat. Neurosci. 8:346-353. Iwanami, A., Kaneko, S., Nakamura, M., Kanemura, Y., Mori, H., Kobayashi, S., Yamasaki, M., Momoshima, S., Ishii, H., Ando, K., Tanioka, Y., Tamaoki, N., Nomura, T., Toyama, Y., and Okano, H. 2005. Transplantation of human

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neural stem cells for spinal cord injury in primates. J. Neurosci. Res. 80:182-190. Kramer, M.F. and Coen, D.M. 2001. Enzymatic amplification of DNA by PCR: Standard procedures and optimization. Curr. Protoc. Mol. Biol. 56:15.1.1-15.1.14. Lindvall, O., Kokaia, Z., and Martinez-Serrano, A. 2004. Stem cell therapy for human neurodegenerative disorders: How to make it work. Nat. Med. 10:S42-S50. Lorenz, W.W., Cormier, M.J., O’Kane, D.J., Hua, D., Escher, A.A., and Szalay, A.A. 1996. Expression of the Renilla reniformis luciferase gene in mammalian cells. J. Biolumin. Chemilumin. 11:31-37. Miyoshi, H., Blomer, U., Takahashi, M., Gage, F.H., and Verma, I.M. 1998. Development of a selfinactivating lentivirus vector. J. Virol. 72:81508157. Naldini, L., Blomer, U., Gallay, P., Ory, D., Mulligan, R., Gage, F.H., Verma, I.M., and Trono, D. 1996. In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263-267. Navarro-Galve, B., Villa, A., Bueno, C., Thompson, L., Johansen, J., and Martinez-Serrano, A. 2005. Gene marking of human neural stem/precursor cells using green fluorescent proteins. J. Gene Med. 7:18-29. Paroo, Z., Bollinger, R.A., Braasch, D.A., Richer, E., Corey, D.R., Antich, P.P., and Mason, R.P. 2004. Validating bioluminescence imaging as a high-throughput, quantitative modality for assessing tumor burden. Mol. Imaging 3:117124. Pichler, A., Prior, J.L., and Piwnica-Worms, D. 2004. Imaging reversal of multidrug resistance in living mice with bioluminescence: MDR1 P-glycoprotein transports coelenterazine. Proc. Natl. Acad. Sci. U.S.A. 101:1702-1707.

Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S., (eds.). 2009. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Rubio, F.J., Bueno, C., Villa, A., Navarro, B., and Martinez-Serrano, A. 2000. Genetically perpetuated human neural stem cells engraft and differentiate into the adult mammalian brain. Mol. Cell Neurosci. 16:1-13. Shah, K., Tung, C.H., Yang, K., Weissleder, R., and Breakefield, X.O. 2004. Inducible release of TRAIL fusion proteins from a proapoptotic form for tumor therapy. Cancer Res. 64:32363242. Shah, K., Bureau, E., Kim, D.E., Yang, K., Tang, Y., Weissleder, R., and Breakefield, X.O. 2005. Glioma therapy and real-time imaging of neural precursor cell migration and tumor regression. Ann. Neurol. 57:34-41. Shah, K., Hingtgen, S., Kasmieh, R., Figueiredo, J.L., Garcia-Garcia, E., Martinez-Serrano, A., Breakefield, X., and Weissleder, R. 2008. Bimodal viral vectors and in vivo imaging reveal the fate of human neural stem cells in experimental glioma model. J. Neurosci. 28:44064413. Snyder, E.Y. and Macklis, J.D. 1995. Multipotent neural progenitor or stem-like cells may be uniquely suited for therapy for some neurodegenerative conditions. Clin. Neurosci. 3:310316. Tang, Y., Shah, K., Messerli, S.M., Snyder, E., Breakefield, X., and Weissleder, R. 2003. In vivo tracking of neural progenitor cell migration to glioblastomas. Hum. Gene Ther. 14:1247-1254. Villa, A., Navarro-Galve, B., Bueno, C., Franco, S., Blasco, M.A., and Martinez-Serrano, A. 2004. Long-term molecular and cellular stability of human neural stem cell lines. Exp. Cell Res. 294:559-570.

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Functional Analysis of Adult Stem Cells Using Cre-Mediated Lineage Tracing

UNIT 5A.2

Diana L. Carlone1 1

Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts

ABSTRACT Lineage-tracing has been used for decades to establish cell fate maps during development. Recently, with the advent of genetic lineage-tracing techniques (employing Cre-lox recombination), it has been possible to permanently mark progenitor/stem cell populations within somatic tissues. In addition, pulse-chase studies have shown that only stem cells are capable of producing labeled progeny after an extensive period of chase. This unit focuses on the protocols used to target putative adult stem cells in vivo. Using these techniques, one should be able to functionally confirm or deny the stem cell capacity of C 2009 by John a given cell population. Curr. Protoc. Stem Cell Biol. 9:5A.2.1-5A.2.15.  Wiley & Sons, Inc. Keywords: tamoxifen-inducible Cre recombination r lineage contribution r reporter activity r whole-mount analysis

INTRODUCTION Adult stem cells are elusive in many tissues. The promise of cell-based therapeutics for the treatment of human disease must first begin with the identification of functionally important stem cell populations. It is generally accepted that stem cells have the capacity for self-renewal and multi-lineage contribution within a given tissue. The Cre-lox system may be used to permanently mark cells of interest so that they can be observed for what they give rise to. If they give rise to no other lineages, they are not stem cells. If they give rise to multiple lineages, and subsequently are shown to self-renew, they are identified as stem cells. This unit focuses on lineage-tracing analysis using tamoxifen-inducible Cre-lox technology (Fig. 5A.2.1) to define the contribution of specific cell populations. This technique has been successfully employed to mark progenitor/stem cell populations in adult tissues. A detailed description of the Cre-lox system can be found elsewhere (Rossant and McMahon, 1999; Nagy, 2000; Branda and Dymecki, 2004). Briefly, Cre recombinase causes recombination of 34-bp loxP sequences and thus deletion of the intervening sequence (Fig. 5A.2.1). It is important to note that the orientation of the loxP sequences with respect to one another determines the recombination outcome (for a review, see Branda and Dymecki, 2004). Both loxP sequences must be in the same orientation for proper excision. Alteration in the orientation results in inversion of the intervening sequence and lack of deletion. While this technology is primarily used to induce tissue-specific knockout of genes in mice, when used to activate reporter genes it can indelibly mark discrete cell populations. The addition of inducible components such as the tamoxifeninducible Cre recombinase (CreER) to the system further allows for the study of temporal relationships between cell populations. Lineage tracing studies typically involve the use of double transgenic mice containing both a Cre-expressing transgene and a Cre reporter transgene. Cells are permanently marked during an initial Pulse and the contribution of their progeny to specific cell lineages is then determined during a period of Chase. Multiple strategies have been Current Protocols in Stem Cell Biology 5A.2.1-5A.2.15 Published online May 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a02s9 C 2009 John Wiley & Sons, Inc. Copyright 

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A IoxP

ATAACTTCGTATAGCATACATTATACGAAGTTAT

B IoxP

gene X

IoxP

Cre recombinase

IoxP

Figure 5A.2.1 Schematic of the Cre-lox system. (A) The sequence for the loxP site is shown. The underlined sequence is the 8-bp core sequence where recombination occurs, and two flanking 13-bp inverted repeats. (B) Schematic of a transgene in which gene X is flanked by two loxP sites. In the presence of Cre recombinase the gene X is deleted leaving only a single loxP.

cell-specific promoter

CreERT X

stop LacZ

OR

Rosa26R reporter

LacZ

hPAP

Z/AP reporter

tamoxifen

LacZ

Cre-Mediated Lineage Tracing

hPAP

Figure 5A.2.2 Schematic of bigenic mouse model systems used for lineage tracing. To perform tracing studies, tamoxifen-inducible CreERT transgenic mice are crossed with either Rosa26R (left) or Z/AP (right) reporter mice. In the absence of ligand, β-galactosidase (LacZ) reporter is not expressed in the Rosa26R mice while Z/AP mice express LacZ. Upon administration of tamoxifen, recombination occurs in a cell-specific manner resulting in either LacZ (left) or human placental alkaline phosphatase (hPAP; right) expression. Once labeled, cells can then be chased to determine their contribution to distinct lineages.

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developed to indelibly mark cells for cell fate mapping. One strategy employs the use of a loxP-flanked dominant transcriptional stop sequence located upstream of a reporter gene. Following Cre-mediated deletion of this stop sequence, constitutive and permanent expression of the reporter gene is induced (see Fig. 5A.2.2; Rosa26R reporter mouse; Soriano, 1999). Another strategy involves the use of tandem reporter genes with the first gene flanked by loxP sites. Under baseline conditions, only the first gene is expressed constitutively. In this scheme, Cre-recombination removes the first gene, allowing the permanent expression of the second (see Fig. 5A.2.2; Z/AP reporter mouse; Lobe et al., 1999). In this unit, the protocols used for analysis of two commonly employed transgenes—β-galactosidase and alkaline phosphatase—are described.

WHOLE-MOUNT ANALYSIS OF β-GALACTOSIDASE ACTIVITY This protocol focuses on whole-mount analysis followed by immunohistochemistry to demonstrate that a discrete population of cells contributes to distinct differentiated lineages. To perform these studies, double transgenic mice containing both a tamoxifeninducible cell-specific CreER transgene and a Cre reporter transgene (β-galactosidase or alkaline phosphatase reporter) are used. As outlined in Critical Parameters, the choice of the Cre transgene as well as the reporter mouse line is dependent upon the scientific question being asked. An example of whole-mount analysis using the β-galactosidase reporter mouse line (Rosa26R) is illustrated in Figure 5A.2.3. While whole-mount analysis has the advantage of allowing for the detection of reporter activity in the context of the intact tissue, it does require subsequent histological analysis to confirm the identity of the marked cells. In this protocol, immunohistochemical analysis is performed using differentiation-specific antibodies to demonstrate the contribution of reporter-positive cells to specific cell lineages. Alternatively, if applicable, histological analysis using specific stains such as periodic acid-Schiff (PAS), which recognizes carbohydrates in tissue sections, can be used to demonstrate histologically that reporter-positive cells are differentiated (see Fig. 5A.2.3C-E).

BASIC PROTOCOL

NOTE: While validation of lineage contribution through immunohistochemical analysis of lacZ-stained regions is described, it is possible to co-label cells fluorescently using both differentiation-specific antibodies and β-galactosidase-specific antibodies using either paraffin or frozen sections.

Materials Tamoxifen-inducible Cre :: Rosa26R or Z/AP bigenic mice Tamoxifen or 4-hydroxytamoxifen (see recipe) Negative control mice (oil-treated bigenic or treated monogenic mice) Positive control mice (Rosa26; Jackson Laboratories cat. no. 002292) Phosphate-buffered saline, Ca++ - and Mg++ -free (CMF-PBS) LacZ fixative, wash, and staining buffers (see recipes) 32% (w/v) paraformaldehyde solution (EMS cat. no. 15714-S) 35% 70%, 80%, 90%, 95%, and 100% ethanol Xylene Paraffin 10 mM sodium citrate, pH 6.0 3% H2 O2 , optional Avidin and biotin blocking solutions (Vector Laboratories cat. no. SP-2001) Normal serum (species selection should match that of the secondary antibody; Sigma) Differentiation-specific antibodies Vectastain ABC Elite kit (species-specific kits are available dependent upon the primary antibody; Vector Laboratories)

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A

B

C

D

E

Figure 5A.2.3 Whole-mount and sectional analysis demonstrate lineage tracing in the small intestine. (A,B) Wholemount analysis of LacZ staining in the intestine following tamoxifen treatment (2-week chase) at low (A; 2×) and high (B; 11.25×) magnification. (C) Frozen sectional analysis of β-galactosidase activity in the small intestine following a 4-week chase. Tissue was subsequently stained with periodic acid-Schiff to detect lineage contribution. Magnification is 40×. (D,E) Histological analysis of lineage contribution by LacZ-positive cells in the small intestine following tamoxifen treatment. Following whole-mount analysis, LacZ-positive regions were paraffin embedded and sections were counterstained with periodic acid-Schiff. Co-labeling of LacZ and periodic acid-Schiff corresponds to goblet (D) and Paneth (E) cells. Magnification is 60×.

DAB substrate kit (Vector Laboratories cat. no. SK-4100) Nuclear Fast Red (Sigma cat. no. N3020) Cytoseal XYL mounting medium (Richard-Allan Scientific cat. no. 8312-4) 1.5-ml microcentrifuge tubes or 6-well tissue culture plates Platform shaker 37◦ C incubator Microtome Microscope slides Coplin jars Pressure cooker, microwave, or water bath Coverslips NOTE: Unless indicated, all steps in this protocol are performed at room temperature.

Induce Cre expression 1. Treat tamoxifen-inducible Cre :: Rosa26R bigenic mice with tamoxifen and control mice with oil. Collect tissue of interest after treatment (pulse) followed by a period of chase. Cre-Mediated Lineage Tracing

Bigenic mice are obtained by crossing tamoxifen-inducible Cre mice and commercially available β-galactosidase reporter Rosa26R mice (Jackson Laboratories cat. no. 003310).

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The dosage, route, and frequency of tamoxifen administration are dependent upon multiple factors, which are outlined in the Critical Parameters section and will need to be empirically determined for each Cre-expressing transgene and tissue of interest. In addition, the appropriate treatment should label single cells that can then be chased to determine their contribution to specific lineages. The length of chase is dependent upon multiple factors as described in Critical Parameters.

2. Wash tissue with ice-cold CMF-PBS to remove any contaminants and place into either a 1.5-ml microcentrifuge tube or 6-well tissue culture plate depending upon the size of tissue to be analyzed. Tissue from oil-treated bigenic or tamoxifen-treated monogenic mice should be used as a negative control to confirm specificity of the reaction. In addition, tissue from the Rosa26 mouse, which constitutively expresses β-galactosidase (Zambrowicz et al., 1997), can be used as a positive control. If necessary, tissues can be cleaned and washed for 5 to 10 min in cold CMF-PBS buffer containing 0.02% (v/v) NP-40 and 0.5 mM DTT prior to fixation. It has been found that this reduces background LacZ staining especially in whole-mount analysis of gastrointestinal tissues. The size/thickness of the tissue can affect the penetration of the staining solution thereby altering the efficiency of labeling. Therefore, using a small tissue biopsy, bisecting the tissue or, if necessary, gently poking holes into the tissue will increase the penetration. If, however, the entire tissue needs to be assayed, then analysis can be performed on tissue sections (see Alternate Protocol 2). Alternatively, if the tissue is thin enough, it can be processed intact while mounted on paraffin in tissue culture dishes using insect pins (Fine Science Tools). This approach is routinely used for whole-mount analysis of small intestine.

Fix tissues 3. Fix tissue in LacZ fixative solution for 1 hr on ice with shaking. To increase penetration and decrease background, the detergent NP-40 (0.02%) can be added to the fixative. Generally, a mild fixative such as glutaraldehyde is used; however, other fixatives can be used, including 4% (w/v) paraformaldehyde, or a combination of fixatives such as 0.2% (v/v) glutaraldehyde/2.0% (v/v) formaldehyde in CMF-PBS. CAUTION: Some fixatives can inhibit or diminish β-galactosidase activity; therefore, it may be necessary to test alternative fixatives.

4. Wash tissue three times with CMF-PBS for 10 min each on ice with shaking.

Stain for LacZ activity 5. Wash with LacZ wash buffer for 10 min on ice with shaking. 6. Incubate tissue in LacZ staining buffer from 1 to 24 hr at 37◦ C in the dark, until a dark color from the substrate reaction is seen, while the background is relatively unstained. The reaction is light sensitive so incubate sample in the dark. Generally, the reaction is stopped within 24 hr. The reaction can also be performed at lower temperatures such as 30o C, which has been shown to reduce background staining. However, a longer incubation time may be necessary.

7. To stop reaction, wash two to three times with CMF-PBS for 20 min each at room temperature with shaking. 8. To preserve tissue and staining, re-fix tissue with 4% paraformaldehyde in CMF-PBS for 1 to 2 hr (longer if necessary) at 4◦ C with shaking. 9. Place tissue in CMF-PBS and store at 4◦ C. Fixed tissue can be stored 3 to 6 months at 4◦ C without diffusion of the LacZ stain.

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Prepare tissues for immunohistochemical confirmation of lineages 10. Dehydrate whole-mount stained tissue through an ethanol series (35%, 70%, 90%, 100%) and xylene. Incubate the tissue two times in each solution, 1 hr each time. Embed in paraffin. The amount of time to dehydrate tissue may vary with the size of the tissue. Paraffin sections are used to confirm the contribution of a marked cell to distinct lineages because they allow better tissue histology than frozen sections.

11. Cut 4-μm sections and mount onto microscope slides. 12. Rehydrate sections through two changes of xylene, 3 min each, followed by two changes of 100% ethanol, 2 min each, and then through an ethanol series (95%, 90%, 80%, and 70%), 1 min each, using Coplin jars. All steps involving histological slides are performed using Coplin jars.

13. Wash with CMF-PBS for 5 min with shaking. For histological analysis of reporter positive cells, paraffin sections after rehydration can be counterstained with Nuclear Fast Red (see below).

Retrieve antigens 14. Perform antigen retrieval by boiling slides in 10 mM sodium citrate, pH 6.0, for 10 min using a pressure cooker, microwave, or water bath. Fixatives such as paraformaldehyde form protein cross-links that may mask antigenic sites giving negative (or weak) immunohistochemical results. Therefore, the antigen retrieval step unmasks the antigens/epitopes in paraffin sections. This buffer is commonly used and works well with most antibodies.

15. Allow slides to cool in buffer for 45 to 60 min. 16. Wash slides two to three with CMF-PBS, 5 min each, with shaking.

Treat to reduce background 17. (Optional) Incubate slides in 3% H2 O2 for 15 min with shaking. This step blocks endogenous peroxidase activity. Because not all tissues exhibit endogenous activity, this step is optional depending upon the tissue of interest.

18. Wash slides three times with CMF-PBS, 5 min each, with shaking. 19. Incubate slides with avidin and biotin blocking solutions, 15 min each, with shaking. Wash three times with CMF-PBS in between each step. Like peroxidase activity, some tissues exhibit endogenous biotin activity; therefore, this step is also optional depending upon the tissue of interest.

20. To reduce non-specific background, block sections with 1% to 5% normal serum in CMF-PBS for 15 to 30 min with shaking. The exact percentage of serum to be used needs to be determined empirically for each antibody. In addition, the normal serum used should be from the same species as the secondary antibody. If necessary, 0.1% to 0.3% Triton X-100 can be used in the blocking solution to further decrease background.

Immunostain slides 21. Incubate sections with primary antibody at appropriate dilution in blocking solution for 1 hr at room temperature to overnight at 4◦ C. The exact incubation conditions for each antibody must be empirically determined. Cre-Mediated Lineage Tracing

22. Wash slides three times with CMF-PBS, 5 min each, with shaking.

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23. Incubate with biotinylated secondary antibody (Vectastain Elite ABC kit) at appropriate dilution in blocking solution per the manufacturer’s instructions for 30 min at 37◦ C. Selection of appropriate Vectastain Elite ABC kit is dependent upon the species used to generate the primary antibody.

24. Wash slides three times with CMF-PBS, 5 min each, with shaking.

Detect antibody binding 25. Incubate sections with ABC reagent 30 min at 37◦ C. 26. Wash slides three times with CMF-PBS, 5 min each, with shaking. 27. Incubate slides in 3,3 -diaminobenzidine (DAB) substrate solution for 1 to 5 min. The DAB solution yields a brown substrate color. Alternatively, the VIP substrate solution (Vector Laboratories cat. no. SK-4600) can be used and yields a purple substrate color. In addition, the precise incubation time must be determined empirically for each antibody.

28. To stop reaction, wash slides with water for 5 min with shaking. 29. If necessary, counterstain sections with Nuclear Fast Red solution for 30 sec to 1 min. This solution is light sensitive. Nuclear Fast Red is diluted 1:1 with distilled water, filtered, and stored for 3 to 6 months at room temperature. This stain can be reused multiple times and re-filtered as needed.

30. Dehydrate slides for 2 min each using 70%, 90%, and 100% ethanol and xylene. 31. Mount slides with coverslips using Cytoseal XYL mounting medium.

WHOLE-MOUNT ANALYSIS OF ALKALINE PHOSPHATASE ACTIVITY Although many researchers use LacZ staining to permanently trace the contribution of stem cells, Cre/lox reporter mice utilizing other reporters such as the enzyme, alkaline phosphatase, can also be used in whole-mount analysis. This protocol can therefore be used as an alternative approach to define the role of a discrete cell population as stem cells using lineage tracing technology.

ALTERNATE PROTOCOL 1

Additional Materials (also see Basic Protocol) Alkaline phosphatase fixative solution (see recipe) AP buffer (see recipe) BM Purple AP substrate (Roche Diagnostics cat. no. 11 442 074 001) PTM buffer (see recipe) 70◦ to 75◦ C incubator Collect tissue 1. Treat tamoxifen-inducible Cre :: Z/AP bigenic mice with tamoxifen and collect tissue at pulse and chase time points. As indicated in the Basic Protocol, tissue from oil-treated bigenic or tamoxifen-treated monogenic mice should be used as a negative control. In addition, constitutively expressing alkaline phosphatase mice can be used as a positive control. If appropriate, alkaline phosphatase and LacZ can be detected in the same tissue. Staining for β-galactosidase must be performed first due to its sensitivity to heat, which is used to reduce endogenous alkaline phosphatase activity (see step 4). After X-gal staining, tissue should be rinsed well with CMF-PBS prior to performing the alkaline phosphatase protocol.

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Fix tissue 2. Fix tissue with alkaline phosphatase fixative solution for 30 min on ice with shaking. 3. Wash tissue three times with CMF-PBS, 10 min each, on ice with shaking. 4. Heat-inactivate endogenous alkaline phosphatase activity by incubating in CMF-PBS 30 min at 70◦ to 75◦ C. 5. Wash with CMF-PBS 10 min at room temperature with shaking.

Detect AP activity 6. Wash with AP buffer 10 min at room temperature with shaking. 7. Stain with BM Purple AP substrate up to 36 hr at 4◦ C, until a dark color from the substrate reaction is seen. Incubation at room temperature will accelerate the reaction but may result in diffusion of the stain.

8. Wash tissue three times with PTM buffer, 10 min each, at room temperature with shaking. 9. To preserve the staining, re-fix the tissue with 4% paraformaldehyde in CMF-PBS for 1 to 2 hr (longer if necessary) at 4◦ C with shaking. 10. Place tissue in CMF-PBS and store for 3 to 6 months at 4◦ C. For immunohistochemical analysis to confirm lineage contribution, tissues can be processed similar to LacZ-stained whole-mount tissue, see Basic Protocol, step 10. ALTERNATE PROTOCOL 2

SECTIONAL ANALYSIS FOR β-GALACTOSIDASE OR ALKALINE PHOSPHATASE ACTIVITY Although whole-mount analysis allows for reporter detection in the intact tissue, analysis of tissue sections for reporter activity identifies the specific marked cell histologically. This protocol can therefore be used as an alternative approach or, in many instances, in combination with whole-mount analysis to further define the role of a discrete cell population as stem cells using lineage tracing technology (see Fig. 5A.2.3).

Additional Materials (also see Basic Protocol) Tissue-Tek OCT (Sakura, cat. no. 4583) 0.2% glutaraldehyde/2 mM MgCl2 in CMF-PBS 0.2% glutaraldehyde in CMF-PBS AP buffer (see recipe) 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium (BCIP/NBT) solution (see recipe or Vector Laboratories cat. no. SK-5400) Cryomolds Cryostat 70◦ to 75◦ C incubator Analyze sections for reporter activity 1. Isolate tissue following chase and pulse time points as determined in whole-mount analysis and place directly into cryomolds containing Tissue-Tek OCT and freeze on dry ice. Alternatively, tissues can be fixed in 4% paraformaldehyde for 1 to 2 hr on ice, incubated in 0.6 M sucrose overnight at 4o C, and then embedded in OCT. Sucrose acts as a cryoprotectant to minimize ice crystal damage, allowing for better microscopic morphology. Cre-Mediated Lineage Tracing

The heating process used during the embedding of tissue for paraffin sections inactivates β-galactosidase; therefore, all sectional analysis is performed using frozen sections.

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2. Section tissue (10-μm), dry up to 2 hr at room temperature, and store at −20◦ C. Stain for β-galactosidase activity 3a. Fix slides in 0.2% glutaraldehyde/2 mM MgCl2 in CMF-PBS for 10 min on ice with shaking. Alternative fixatives such as 4% paraformaldehyde may be used; however, as indicated with the whole-mount analysis, reporter activity may vary with harsher fixatives.

4a. Wash slides two to three times with LacZ wash buffer, 10 min each, at room temperature with shaking. 5a. Incubate in LacZ staining buffer 1 to 24 hr at 37◦ C in the dark. 6a. To stop the reaction, wash slides three times with CMF-PBS, 10 min each, followed by a quick rinse in water at room temperature. 7a. Counterstain sections with Nuclear Fast Red solution for 30 sec to 1 min. 8a. Dehydrate slides for 2 min each using 70%, 90%, and 100% ethanol and xylene. 9a. Mount slides with coverslips using Cytoseal XYL mounting medium. Stain sections for alkaline phosphatase activity 3b. Fix slides in 0.2% glutaraldehyde in CMF-PBS for 10 min on ice with shaking. 4b. Wash slides three times with CMF-PBS, 5 min each, at room temperature with shaking. 5b. Inactivate endogenous alkaline phosphatase by incubating slides in CMF-PBS for 30 min at 70◦ to 75◦ C. 6b. Wash slides with CMF-PBS for 10 min at room temperature with shaking. 7b. Wash slides with AP buffer for 10 min at room temperature with shaking. 8b. Stain slides with BCIP/NBT solution for 10 to 30 min at room temperature. Staining solution should be placed directly onto the slides. Sensitivity of substrate can be increased by lengthening the incubation time.

9b. Wash slides in CMF-PBS and process as indicated above in steps 7a through 9a.

REAGENTS AND SOLUTIONS For all solutions, use deionized, distilled water or equivalent in recipes and protocol steps. Unless indicated, all solutions are made up in water. Suppliers for non-common chemicals are indicated.

Alkaline phosphatase fixative solution 0.2% (v/v) glutaraldehyde 50 mM EGTA, pH 7.3 100 mM MgCl2 0.02% (v/v) NP-40 0.01% (w/v) sodium deoxycholate Prepare fresh To increase the penetration of alkaline phosphatase (AP) substrates, 0.02% NP-40 and 0.01% sodium deoxycholate have been added to the fixative.

AP buffer 10 mM Tris·Cl, pH 9.5 100 mM NaCl continued

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10 mM MgCl2 Store up to 1 year at room temperature LacZ fixative solution 0.2% (v/v) glutaraldehyde 2 mM MgCl2 CMF-PBS Prepare fresh LacZ staining buffer 0.5 mg/ml X-gal (stock is 40 mg/ml in dimethylformamide) 5 mM K3 Fe(CN)6 5 mM K4 Fe (CN)6 -3H2 O LacZ wash buffer (see recipe) Prepare fresh Staining buffer should be made fresh each time. Use CMF-PBS at approximately pH 7.4 for all steps. If necessary, use CMF-PBS at pH 8.0 for LacZ staining buffer to decrease background β-galactosidase activity.

LacZ wash buffer 2 mM MgCl2 0.01% (w/v) deoxycholate 0.02% (v/v) NP-40 CMF-PBS Store up to 1 year at room temperature NBT/BCIP stain solution 100 mM Tris·Cl, pH 9.5 100 mM NaCl 50 mM MgCl2 0.01% (w/v) sodium deoxycholate 0.02% (v/v) NP-40 337 μg/ml nitroblue tetrazolium salt (NBT; Sigma cat. no. N6876) 175 μg/ml 5-bromo-4-chloro-3-indolyl phosphate, disodium salt (BCIP; Sigma cat. no. B1026) Prepare fresh PTM buffer 0.1% (v/v) Tween-20 2 mM MgCl2 CMF-PBS Store up to 1 year at room temperature Tamoxifen or 4-hydroxytamoxifen Resuspend tamoxifen (Sigma cat. no. T5648) or 4-hydroxytamoxifen (70% Z isomer, 30% E isomer 4-OHT; Sigma cat. no. H6278) at a concentration of 10 to 20 mg/ml in peanut oil (Indra et al., 1999; K¨uhbandner et al., 2000). Add 500 μl of 100% ethanol to 100 mg of tamoxifen followed by 9.5 ml of peanut oil. Dispense into aliquots and store for up to 4 weeks at −20◦ C. Corn oil can also be used to resuspend tamoxifen or 4-OHT. Cre-Mediated Lineage Tracing

Tamoxifen is fairly soluble while 4-OHT requires sonication or heating. Both will precipitate at cold temperatures; therefore, it may be necessary to heat or resonicate prior to use.

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COMMENTARY Background Information The identification of tissue progenitor/stem cells requires demonstrating both their selfrenewal potential and their capacity to contribute to multiple cell lineages within a tissue. While current studies use genetic models to experimentally demonstrate stem cell function, the scientific literature is full of alternate and historical approaches to map the fate of a cell (reviewed in Stern and Fraser, 2001). Early studies focused on deciphering cell lineage using a variety of techniques to follow cells and their descendants. For example, endogenous expression of alkaline phosphatase by germ cells allowed for tracing of these cells throughout development (Chiquoine, 1954). In addition, direct visualization by the use of pigmentation differences among cells has allowed for the construction of lineage trees in developing organisms, including the complete cell fate map of the nematode, Caenorhabditis elegans (Sulston et al., 1983; Thomas et al., 1996). This technique is limited however by an inability to identify and trace single cells over time and the problems inherent to increasingly opaque embryos. To combat these problems, a variety of approaches have been taken to mark cells. The use of vital dyes to trace living cells was attempted but was found to be ineffective due to their water solubility, which resulted in transfer of dye to unrelated cells. Eventually, multicolored carbocyanine dyes were generated, which are lipid soluble/water insoluble and localize within the cell membrane (Axelrod, 1979; Serbedzija et al., 1989; YablonkaReuveni, 1989; Eagleson and Harris, 1990). While cells readily take up these dyes, it proved difficult to mark single cells, thus this technique has been used to track the fate of cell populations. Additional strategies have included the use of radiolabeled compounds to mark cells prior to introduction into embryos as well as the generation of interspecies chimeras (e.g., chick/quail), which relies upon species-specific differences in cell pigmentation and size to distinguish between donor and host cells (Le Douarin, 1973; Dupin et al., 1998). Marking and tracing of single cells in both vertebrates and invertebrates have been accomplished by single-cell injection with inert tracers such as the enzyme horseradish peroxidase, or fluorescently-labeled compounds such as dextran or lysine (Weisblat et al., 1978; Lawson et al., 1986; Gimlich and Braun, 1985;

Peralta and Denaro, 2003). While technically challenging, very elegant cell fate mapping is possible with such approaches. The biggest disadvantage is that the tracer becomes diluted in dividing cells. To determine the fate of cells that might migrate during development, colloidal goldlabeled monoclonal antibodies were developed and used to track the descendant of cells that express a common surface antigen regardless of their position in the embryo (Stern and Canning, 1990). Antigen-expressing cells internalize the antibody/gold complex and pass it on to their descendants. However, the marker can only be detected for a few divisions. To overcome this problem, several groups developed retroviral vectors that would label cells through the introduction of marker genes, e.g., alkaline phosphatase or β-galactosidase (Cepko et al., 1984; Sanes et al., 1986). Diluted viral stock solutions, as well as replicationdeficient strains, were subsequently employed to increase the probability that marked cells would be clonally derived. This approach has been used to trace cell lineages in the nervous system (Cepko et al., 1984; Price et al., 1987). Given that viral targeting is not celltype-specific, these lineage-mapping studies must be performed retrospectively. In addition, it is impossible to rule out that adjacent cells were not also initially labeled via an independent infection event. To address this, complex retroviral libraries were generated (Golden et al., 1995) with the belief that the infection was a random event and thus it would be highly unlikely that adjacent cells would be infected by the same retrovirus. Distinction between the various retroviruses could be confirmed by PCR. Although this approach increased the rigor, the analysis remained retrospective and required that each cell type be analyzed by PCR to determine their relatedness to neighboring labeled cells. Therefore, this method required the ability to isolate the cell types of interest. Alternatively, spontaneous DNA recombination events (+/− mutagens) have been used to retrospectively trace cells via activation of marker expression in transgenic mice (Bonnerot and Nicolas, 1993; Bjerknes and Cheng, 1999). Once labeled, cells are transplanted into adult animals to demonstrate cell fate mapping. The contribution of a putative stem cell to various cell lineages is assessed through analysis of marker expression or, alternatively, identification of the Y chromosome when

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male donor cells were injected into female recipients. This approach has been used to study hematopoietic stem engraftment and multi-lineage contribution. While these approaches have yielded important insight into cell fate, increasingly, the use of site-specific recombinase systems such as Cre-lox has become the gold standard for cell lineage studies. This technology allows for the control over the temporal and spatial marking of selective cell populations through the use of gene-specific promoters to regulate recombinase expression (for review see Sauer, 1998; Nagy, 2000; Branda and Dymecki, 2004). To regulate the onset of Cre expression independent from endogenous regulatory elements, inducible Cre recombinases have been generated that allow for refined labeling of specific single cells in response to ligand administration (Furth et al., 1994; Feil et al., 1996; Kellendonk et al., 1996; Rivera et al., 1996; Brocard et al., 1998; Danielian et al., 1998; Utomo et al., 1999; Sch¨onig et al., 2002). Specifically, in the tamoxifen-inducible Cre-lox system (used in this unit), Cre recombinase (CreER) is fused to a mutated ligand-binding domain of the estrogen receptor, which specifically binds tamoxifen and not the endogenous estrogen. In the absence of ligand, CreER is retained within the cytoplasm. Upon tamoxifen administration, the CreER protein translocates to the nucleus where it excises the loxP-targeted site. Furthermore, the commercial availability of a variety of Cre-reporter mouse lines, which upon recombination express colorimetric, enzymatic, or fluorescent proteins, allows for greater diversity in the use of lineage-tracing to define stem cells.

Critical Parameters and Troubleshooting

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While the lineage-tracing approach can demonstrate “stemness” of a particular cell population, it does require some a priori knowledge. First, one must determine whether a specific gene selectively marks the cell of interest. Second, determine whether a Cre mouse line already exists in which the promoter of the gene of interest regulates Cre recombinase expression. Preferably this line should be inducible, allowing for controlled temporal marking of the cell. Third, determine the Cre reporter mouse line to be used. The choice of which Cre reporter line to use is dependent upon several things including whether the reporter is expressed in the cell and tissue of interest as well as the type of analysis that

will be performed. Although most reporters are considered to be ubiquitously expressed, it has been found that is not always the case; therefore, when possible, confirming the expression of the reporter in the cell of interest is optimal. In addition, while this unit focuses on β-galactosidase and alkaline phosphatase reporters, fluorescent Cre reporter mouse lines have been used to trace lineage contribution. Generally, colorimetric and enzymatic reporters are convenient to use for confirming lineage contribution by a stem cell; however, fluorescent reporters have the added advantage of allowing for single-cell isolation and analysis via flow cytometry. The following Websites contain additional information on the various Cre and Cre reporter mouse lines available for lineage tracing: The Jackson Laboratory: http://www.jax.org or Dr. Andras Nagy’s laboratory Website: http://www.mshri.on.ca/nagy. Although stem cells have been identified in tissues that are highly regenerative such as the skin, intestine, and blood, adult stem cells in other tissues may require some type of regenerative stimulus such as injury to awaken them. Therefore, prior knowledge of the mechanism(s) involved in stem cell activation within the tissue of interest will be advantageous before performing lineage tracing studies. Administration of the inducing agent varies depending upon the model system used. In the case of the tamoxifen-inducible Cre mouse models, tamoxifen can be administration intraperitoneally (i.p.), subcutaneously (s.c.), and orally (p.o.) as well as by pellet implants. The amount and length of administration is highly dependent upon the strength and expression pattern of the promoter regulating the Cre recombinase and will have to be determined empirically by each researcher. Tamoxifen is generally given at a dose of 1 to 2 mg/day i.p. for up to 5 days. Analysis of reporter activity is determined at the end of the pulse period (1 to 3 days after the last injection) at which time cell-specific labeling should be detected. It is important to note that this regimen should be used as a starting point and modifications in either length of delivery and/or dosage may be necessary depending upon the tissue of interest. If weak or no labeling is detected, then increasing the dose of tamoxifen may be warranted. Alternatively, 4-hydroxytamoxifen (4-OHT), a metabolite of tamoxifen, which exhibits a more potent activity, may be used instead to increase labeling. However, tamoxifen is often preferred over 4OHT because it is more soluble in solution and less costly. It is worth noting that high

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levels of tamoxifen or 4-OHT administered i.p. can have deleterious affects on mice. Varying the timing or mode of administration can circumvent this problem, e.g., administering i.p. injections every other day instead of daily or injecting s.c. or p.o. instead of i.p. Once labeled, the time period in which to analyze the cell’s contribution to various progeny (chase) must be determined. The exact time is dependent upon the tissue of interest as well as whether the stem cell population requires a regenerative stimulus to induce new lineage development. For a general reference, the chase can range from weeks to months. As indicated in the Basic Protocol, contribution of the marked stem cell to differentiated progeny can be confirmed by co-localization of the reporter with differentiation-specific markers by immunohistochemistry or immunofluorescence (depending upon the reporter). This approach however relies upon the availability of specific antibodies. If none are available, alternative approaches may be used such as differentiation-specific gene marker analysis of isolated reporter-positive cells using flow cytometry. As indicated earlier, multiple fluorescent reporter mouse lines are available as well as commercially available flow cytometric kits for β-galactosidase activity (Invitrogen). High background LacZ staining may be due to endogenous enzymatic activity, which has been reported in a variety of tissues. Endogenous β-galactosidase is normally active at low pH (∼4) while bacterial β-galactosidase is active at a more neutral pH. Therefore, increasing the pH of CMF-PBS in the LacZ wash and staining buffers should decrease background LacZ staining. CMF-PBS at a pH between 7.4 and 8.0 is usually used. Extreme alkaline conditions (pH 8.5 to 9.0), however, can inhibit bacterial β-galactosidase activity. In addition, the incubation temperature and time can affect background staining. Minimizing the reaction time to 2 hr and reducing the temperature from 37◦ C to room temperature dramatically reduces background staining. It is important to note that the optimal staining conditions for each tissue needs to be determined empirically using both positive and negative control tissues. There are several possible explanations for a lack of or low reporter activity. As a general rule, tissues from positive control mice are used as a control to validate that the assay is functioning correctly. Absence of reporter activity may be due to either the re-

porter or the Cre recombinase not being expressed in the cell type of interest. To confirm the cellular specificity of Cre recombinase expression, in situ hybridization or immunohistochemical analysis may be performed. As indicated above, although most reporters are believed to be globally expressed upon recombination, variegated expression both across tissues and within select tissues has been found. Therefore, confirmation of reporter expression in the cell and tissue of interest is essential before performing lineage tracing experiments. This can be done by assessing reporter activity in a positive control mouse such as Rosa26 (Zambrowicz et al., 1997), which expresses β-galactosidase from the same genomic locus as the Cre reporter mouse line, suggesting comparable control elements. Alternatively, decreased penetration may affect the staining efficiency in whole-mount analysis. This can be addressed by either analyzing smaller tissue pieces, addition of detergents, or alternatively performing sectional analysis. Finally, low levels of reporter activity may be due to low efficiency of recombination. The strength of the promoter and the number of cells expressing the gene of interest can affect the efficiency of labeling. Increasing the dosage and/or altering the administration regimen may yield higher recombination efficiencies. Alternatively, it may be necessary to use a different inducible system such as the doxycycline system (Sch¨onig et al., 2002) to increase the relative efficiency.

Anticipated Results In response to tamoxifen administration, reporter activity should be detected in a small population of cells preferably single cells at the pulse time point. If multiple cells are labeled, then decreasing the tamoxifen dosage or length of administration should yield single cells. Depending upon the tissue, wholemount analysis of single cells may be difficult; therefore, confirmation of single cells may require sectional analysis. Following this, if a cell functions as a stem cell, then contribution of label to other cell types including differentiated cells within a defined time period should be observed.

Time Considerations The time period required to detect lineage contribution depends upon a number of factors including the regenerative nature of the tissue. For example, the intestine is highly regenerative and turns over every 4 to 5 days; therefore,

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it would require a relatively short chase period to detect stem cells. Alternatively, a more quiescent stem cell population may take much longer to be activated and would therefore require an extended time period before lineage contribution would be detected. In addition, it is important to note that the requirement of inducing regeneration by tissue injury (thereby activating a stem cell population) can further extend the chase time period. Thus, a general time-frame for conducting these experiments relies upon the tissue of interest and will have to be determined by each researcher. It may take a chase weeks to months to detect a stem cell contribution.

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Furth, P.A., St. Onge, L., B¨oger, H., Gruss, P., Gossen, M., Kistner, A., Bujard, H., and Hennighausen, L. 1994. Temporal control of gene expression in transgenic mice by a tetracycline-responsive promoter. Proc. Natl. Acad. Sci. U.S.A. 91:9302-9306.

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Weisblat, D.A., Sawyer, R.T., and Stent, G.S. 1978. Cell lineage analysis by intracellular injection of a tracer enzyme. Science 202:1295-1298.

Soriano, P. 1999. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat. Genet. 21:70-71. Stern, C.D. and Canning, D.R. 1990. Origin of cells giving rise to mesoderm and endoderm in chick embryo. Nature 343:273-275. Stern, C.D. and Fraser, S.E. 2001. Tracing the lineage of tracing cell lineages. Nat. Cell Biol. 3:E216-E218. Sulston, J.E., Schierenberg, E., White, J.G., and Thomson, J.N. 1983. The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev. Biol. 100:64-119.

Yablonka-Reuveni, Z. 1989. The emergence of the endothelial cell lineage in the chick embryo can be detected by uptake of acetylated low density lipoprotein and the presence of a von Willebrand-like factor. Dev. Biol. 132:230240. Zambrowicz, B.P., Imamoto, A., Fiering, S., Herzenberg, L.A., Ker, W.G., and Soriano, P. 1997. Disruption of overlapping transcripts in the ROSA βgeo 26 gene trap strain leads to widespread expression of β-galactosidase in mouse embryos and hematopoietic cells. Proc. Natl. Acad. Sci. U.S.A. 94:3789-3794.

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Magnetic Resonance Imaging of Human Embryonic Stem Cells

UNIT 5A.3

Jaehoon Chung,1 Mayumi Yamada,1 and Phillip C. Yang1 1

Stanford University School of Medicine, Stanford, California

ABSTRACT Magnetic resonance imaging (MRI) may emerge as an ideal non-invasive imaging modality to monitor stem cell therapy in the failing heart. This imaging modality generates any arbitrary tomographic view at high spatial and temporal resolution with exquisite intrinsic tissue contrast. This capability enables robust evaluation of both the cardiac anatomy and function. Traditionally, superparamagnetic iron oxide nanoparticle (SPIO) has been widely used for cellular MRI due to SPIO’s ability to enhance sensitivity of MRI by inducing remarkable hypointense, negative signal, “blooming effect” on T2*-weighted MRI acquisition. Recently, manganese chloride (MnCl2 ) has been reported by our laboratory for its ability as a contrast agent to track biological activity of viable cells. Hyperintense, positive signals can be achieved from the Mn2+ -labeled stem cells on T1-weighted MRI acquisition. Cytotoxicity is a potential drawback of Mn2+ labeling of the cells. However, in our laboratory the labeling method has been optimized to minimize cytotoxic effects. This article describes two different magnetic labeling methods of human embryonic stem cells (hESC) using SPIO and MnCl2 . Curr. C 2009 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 10:5A.3.1-5A.3.9.  Keywords: human embryonic stem cell (hESC) r magnetic resonance imaging (MRI) r superparamagnetic iron oxide (SPIO) r manganese

INTRODUCTION Magnetic resonance imaging (MRI) may emerge as an ideal non-invasive imaging modality to monitor stem cell therapy in the failing heart. This imaging modality generates any arbitrary tomographic view at high spatial and temporal resolution with exquisite intrinsic tissue contrast. This capability enables robust evaluation of both the cardiac anatomy and function. Traditionally, superparamagnetic iron oxide nanoparticle (SPIO) has been widely used for cellular MRI due to SPIO’s ability to enhance sensitivity of MRI by inducing remarkable hypointense, negative signal, “blooming effect” on T2*-weighted MRI acquisition (Fig. 5A.3.1). Recently, manganese chloride (MnCl2 ) has been reported by our laboratory for its ability as a contrast agent to track biological activity of viable cells. Hyperintense, positive signals could be achieved from the Mn2+ -labeled stem cells on T1-weighted MRI acquisition (Fig. 5A.3.2). Cytotoxicity was a potential drawback of Mn2+ labeling of the cells but in our laboratory, the labeling method has been optimized to minimize cytotoxic effects. This article describes two different magnetic labeling methods of human embryonic stem cells (hESC) using SPIO (Basic Protocol) and MnCl2 (Alternate Protocol). NOTE: All procedures must be performed in a sterile cell culture hood. All solutions and equipment in contact with live cells must be sterile, and aseptic technique must be used accordingly. NOTE: All culture incubations must be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Genetic Manipulation of Stem Cells Current Protocols in Stem Cell Biology 5A.3.1-5A.3.9 Published online August 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a03s10 C 2009 John Wiley & Sons, Inc. Copyright 

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Figure 5A.3.1 Magnetic resonance imaging of SPIO-labeled human embryonic stem cells at a coronal view. The different quantities of SPIO-labeled hESC (a) 2 million, (b) 1 million, (c) 0.5 million, (d) 0.1 million, (e) 0.05 million, and control (non-labeled designated with an *).

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Figure 5A.3.2 3 × 106 MnCl2 -labeled hESC at an axial view using the following concentrations: (i) control [0.9% (w/v) sodium chloride solution only], (ii) 0.01 mM, (iii) 0.05 mM, (iv) 0.10 mM, (v) 0.50 mM, (vi) 1.00 mM, and (vii) 3.00 mM. Dose-appropriate increase in T1-weighted positive contrast is seen up to 1.00 mM. hESC are indicated by a black arrow.

BASIC PROTOCOL

DIRECT MAGNETIC LABELING OF HUMAN EMBRYONIC STEM CELLS (hESC) USING SPIO Direct labeling of hESC with SPIO is a convenient and robust method. This technique requires the addition of transfection agents such as protamine sulfate, poly-L-lysine (PLL) or electroporation to increase efficiency of labeling (Frank et al., 2002; Suzuki et al., 2007). Here we introduce a protocol using protamine sulfate (PS). The dextrancoated SPIO particles carry a negative charge. By coating SPIO with positively charged PS, neutral or slightly positive electrical charge can be formed on the surface of the SPIO-PS complex. Consequently, electrostatic interaction between the cell membrane and SPIO-PS complex enhances cellular uptake of SPIO-PS complex via mechanisms including endocytosis, membrane disruption, or passive diffusion.

Materials Magnetic Resonance Imaging of hESCs

H9 hESC (WiCell) hESC medium (see recipe) SPIO (Feridex, Bayer Healthcare Pharmaceuticals)

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Protamine sulfate (PS, American Pharmaceutical Partners) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) Heparin sodium (Fujisawa) Phantom (see recipe) Matrigel-coated 100-mm tissue culture dishes (BD Biosciences; coat dishes according to manufacturer’s instructions) 0.2-ml PCR tubes Signa 3T Excite HD scanner (GE Health System) and an array knee coil with GRE or SPGR sequence 1. Two days prior to the experiment, seed hESC onto Matrigel-coated 100-mm tissue culture dishes in 10 ml hESC medium. This procedure will allow removal of MEF and recovery of hESC culture.

2. One day prior to the experiment, dilute SPIO with hESC culture medium at a final concentration of 50 μg/ml. 3. Add clinical grade PS to the SPIO-containing medium to a final concentration of 6 μg/ml. Shake the mixture vigorously for 5 to 10 min. This procedure will enhance coating of SPIO with PS, generating SPIO-PS complex.

4. Incubate hESC with SPIO-containing medium overnight (8 to 12 hr) in the incubator. This incubation time varies depending on the cell types. We could achieve satisfactory labeling of hESC, human mesenchymal stem cells (hMSC), and mouse embryonic stem cells (mESC) after an 8-hr incubation.

5. Wash hESC two times, each time with 15 ml CMF-PBS. 6. Dilute heparin sodium with CMF-PBS to a final concentration of 10 U/ml. Wash hESC once with 15 ml heparin sodium–containing CMF-PBS. This procedure will allow elimination of SPIO-PS complex on the extracellular surface.

7. Suspend 2 × 106 hESC in 200 μl of CMF-PBS and transfer cell pellet to a 0.2-ml PCR tube. For the negative control, put the same number of non-labeled hESC in one PCR tube. The SPIO-labeled cell pellet will look dark-brown in color (Fig. 5A.3.3).

8. Place the PCR tubes onto the phantom gently to make sure no air is trapped in between the PCR tubes and the phantom to minimize any artifact from the air (Fig. 5A.3.3). The phantom will stabilize the PCR tubes and will prevent artifacts from the surrounding air. Air can induce hypointense signal on T2-weighted sequences. If a crack is noted on the phantom, it can be filled up with water to prevent artifacts.

9. Scan the hESC using T2-weighted sequences. We scan cells using Signa 3T Excite HD scanner and an array knee coil with GRE or SPGR sequence and the following parameters: TR (repetition time) = 100 to 200 msec, TE (echo time) = 20 to 30 msec, FOV (field of view) = 12 × 12 cm, and matrix = 192 × 192, NEX 1. MRI parameters should be optimized for different cell types, SPIO concentration, and magnetic field strength. For in vivo experiments, follow the same labeling method from steps 1 to 6 above. Then, transplant the optimal number of cells in the organ of interest. In our laboratory, 0.5 × 106 SPIO-labeled hESC are injected into the mouse heart for the myocardial infarction model.

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Figure 5A.3.3 Phantom for cellular imaging. The phantom is solidified within a plastic container. Any size plastic container can be used where PCR tubes can be placed stably. PCR tubes containing SPIO-labeled hESC cell pellets were placed onto the phantom. SPIO-labeled cells have a brown color (red arrows) and negative control, non-labeled cells look whitish (black arrow).

ALTERNATE PROTOCOL

DIRECT LABELING OF HUMAN EMBRYONIC STEM CELLS USING MANGANESE CHLORIDE SPIO has been employed to track and localize the transplanted stem cells with high sensitivity. However, this method does not monitor the viability of transplanted stem cells (Kraitchman et al., 2003). On the other hand, MnCl2 has been known to enter viable cells via voltage-gated calcium (Ca2+ ) channels. When the cells are biologically active, MnCl2 accumulates intracellularly to generate a T1-shortening effect to induce a hyperintense, bright signal on T1-weighted MRI acquisition (Aoki et al., 2006). The following protocol is for direct labeling of hESC using manganese and a T1-weighted MRI sequence.

Materials H9 hESC (WiCell) hESC medium (see recipe) TrypLE express (Invitrogen) MnCl2 (Sigma) 0.9% (w/v) sodium chloride (9 mg sodium chloride/ml of distilled water) Phantom (see recipe) Matrigel-coated 100-mm tissue culture dishes (BD Biosciences; coat dishes according to manufacturer’s instructions) Hemacytometer 15-ml conical tubes 0.2-ml PCR tubes Signa 3T Excite HD scanner (GE Health System) Magnetic Resonance Imaging of hESCs

1. Two days prior to the experiment, seed hESC onto Matrigel-coated 100-mm tissue culture dishes in 10 ml hESC medium.

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2. On the day of experiment, remove the culture medium from the dish. Add 1.5 ml of TrypLE express and incubate the cells 5 min. Add 10 ml of culture medium and dissociate the cells by pipetting up and down several times. Wash the cells by centrifuging 5 min at 100 × g, room temperature. Resuspend the cells in culture medium and count cells using a hemacytometer. 3. Aliquot 3 × 106 hESC into one 15-ml conical tube. 4. Dissolve fresh MnCl2 with 0.9% sodium chloride solution to make a 0.1 mM MnCl2 solution. IMPORTANT NOTE: Always make a fresh MnCl2 solution. MnCl2 is easily degraded in solution.

5. Incubate the cells with 5 ml 0.1 mM MnCl2 for 30 min. Incubate negative control cells in 5 ml 0.9% sodium chloride alone. 6. Centrifuge the cells 5 min at 100 × g, room temperature. Aspirate the supernatant and wash the labeled cells twice, each time with 15 ml CMF-PBS. 7. Suspend each pellet in 200 μl CMF-PBS and transfer into 0.2-ml PCR tubes. Manganese-labeled pellets are usually white. For in vivo experiments, transplant these labeled cells in an organ of interest.

8. For an in vitro experiment, place the PCR tubes onto the phantom as described above (step 8 from the Basic Protocol). 9. Perform MRI scanning with T1-weighted spin echo sequence with the following parameters: TR; Repetition time = 800 msec. TE; Echo time = minimum, FOV; field of view = 12 × 12 cm; matrix = 192 × 192, NEX 1. MRI parameters should be optimized for the contrast effect depending on the cell type and magnetic field. Comparable magnetic labeling efficiency has been achieved with different hESC media such as DMEM/F-12 (Invitrogen) or mTeSR (STEMCELL Technologies).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

Human embryonic stem cell medium (500 ml) 400 ml Knockout DMEM (Invitrogen) 100 ml Knockout serum replacement (Invitrogen) 5 ml of 10 mM non-essential amino acids (Invitrogen) 5 ml of 200 mM L-glutamine (Invitrogen) 3.5 μl of 14.3 M 2-mercaptoethanol (Sigma) 10 μg/ml recombinant human bFGF (R&D system) Store up to 1 week at 4◦ C Phantom 0.7% (w/v) agar (Sigma) 1% (w/v) copper sulfate (Sigma) Make solution with distilled water and microwave at medium high for 5 min Solidify in any plastic container before usage (Fig. 5A.3.3) Store up to 6 months at room temperature

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Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) 0.144 g KH2 PO4 9.0 g NaCl 0.795 g Na2 HPO4 ·7H2 O H2 O to 1 liter Adjust to pH 7.4, if necessary Store indefinitely at room temperature COMMENTARY Background Information

Magnetic Resonance Imaging of hESCs

Magnetic resonance imaging (MRI) excites hydrogen protons, using a computerized program called the pulse sequence, to acquire and process the signal released from the excited protons. A pulse sequence consists of radiofrequency and gradient pulses which are carefully controlled in duration and timing to generate images of interest. In MRI, the critical properties are proton density and two basic relaxation times described as spin-lattice and spin-spin relaxation times denoted as T1 and T2, respectively. Relaxation time refers to the time required for the excited tissue to return to the equilibrium state after a radiofrequency pulse is applied. T1 and T2 depend on the proton density of each tissue. Fluid has longer T1 while fat has a shorter T1. T2 is usually shorter than T1 for a given tissue. Fluid has longer T2 while fat has a shorter T2. T2* refers to the effect of additional field inhomogeneity, which contributes to the dephasing signal. T2* is usually shorter than T2. In general, tissues with a long T2 give high signal intensities in T2-weighted images while a long T1 generates a weak signal. Exquisite intrinsic contrast achieved in MRI due to the differences in these relaxivity properties generates detailed images of the anatomy and morphology. Deeply located tissues in small animals can be visualized with high sensitivity by an MRI system. The location and number of the receive coil, the configuration of the coil element, and the magnetic field strength all play important roles. However, the ability of MRI to acquire images from any arbitrary tomographic plane enables detection of the cells in small animals with high sensitivity. In our laboratory, we image SPIO-labeled hESC in an 8-week-old SCID mouse using a 3.0 Tesla clinical scanner (Fig. 5A.3.4). Superparamagnetic iron oxide nanoparticles (size ∼100 nm) can induce a strong magnetic field inhomogeneity (dephasing signal) in the hydrogen atoms of water molecules during magnetic resonance imaging. When SPIO are taken up by the cells, the nanopar-

ticles create significant dephasing of protons, which consequently reduce T2* relaxation times. These properties enable robust visualization of SPIO-labeled cells through strong hypointense, negative signals described as the blooming effect (Bulte et al., 2001). Our data show significant contrast by in vitro MRI 14 days after labeling hESC with SPIO. In vivo MRI showed SPIO-induced contrast 20 days following transplantation of SPIOlabeled hESC in the mouse heart. Despite high sensitivity in the detection of the cells in the range of 10-9 mole/liter, this blooming effect may produce a large signal void at the region of interest to confound the MRI signal from the surrounding artifact and corrupt the anatomical details or the physiological function of the target tissue. Moreover, in vivo experiments have demonstrated that SPIO-labeled cells provide high sensitivity to detect the anatomical location of the cells. However, SPIO labeling does not provide any biologic information such as the viability of transplanted cells because of the non-specific uptake by the macrophages of the residual SPIO particles in the surrounding tissue from dead SPIO-labeled cells (Chen et al., 2008; Li et al., 2008). To address these limitations, our laboratory developed the Mn2+ labeling protocol for stem cells. Mn2+ is transported into the cellular cytoplasm of biologically active cells through a voltage-gated Ca2+ channel. These channels have high affinity for Ca2+ and its analog, such as Mn2+ , to accumulate within the cytoplasm by binding to specific sites on nucleic acid and intracellular proteins. Intracellular Mn2+ induces a T1-shortening effect, which allows clear delineation of the cells of interest with hyperintense, positive signal (Lin and Koretsky, 1997). Therefore, this contrast mechanism enables correlation between cellular viability and a T1-weighted positive signal. Our data shows Mn2+ -induced contrast effect is noted for 4 to 5 days after in vivo labeling (Fig. 5A.3.5). After these cells die, Mn2+ diffuses passively out of these dead cells. Consequently, decreased concentration of Mn2+

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Figure 5A.3.4 In vivo MRI of hESC transplanted into the murine myocardium. Following direct labeling of hESC with SPIO and protamine sulfate, 0.5 × 106 hESC were transplanted into the left ventricle as indicated by negative dephasing signal (black arrow). This mouse was scanned using a 3T clinical MRI scanner (GE Healthcare System).

results in reduced T1-shortening effect and the contrast effect is lost. Currently, two distinct classes of iron oxide particles are available based on the hydrodynamic particle size. The mean diameter of SPIO is ∼100 nm. Ultrasmall superparamagnetic iron oxide particles (USPIO) are ∼40 to 50 nm. Both nanoparticles have similar chemical structures consisting of dextran coating to prevent destabilization and agglomeration of the colloidal suspension to enhance solubility in aqueous or biological media. Because of this chemical structure, both of these agents are biocompatible and SPIO is FDA-approved for imaging of liver lesions. Our studies have shown that mouse and human embryonic stem cell viability and differentiation capacity are not altered with SPIO labeling. However, other studies reported in vitro differentiation capacity of mesenchymal stem cells into chondrocyte lineage was reduced after SPIO labeling in a dose-dependent manner, whereas osteogenic and adipogenic differentiation was intact (Kostura et al., 2004). Direct SPIO labeling of hESC is simple and straightforward. Higher efficiency of iron-oxide labeling is achieved by adding

transfection agents such as PLL, PS or lipofectamine. All these transfection agents neutralize the negatively charged SPIO to facilitate the attraction and binding of slightly positive or neutral complex to the negatively charged cell membrane. The mechanisms by which these complexes enter the cell have not been completely investigated but they probably include endocytosis, invagination, or diffusion. Similarly, MnCl2 is a simple, robust, and safe method to label hESC. Mn2+ is an essential trace element in the human body with electrochemical properties analogous to Ca2+ . Using the voltage-gated Ca2+ channels, not only are the hESC localized with T1-weighted, positive signal, but the biological properties of the cells can also be determined. Nevertheless, the sensitivity/ability to detect cells is still higher with SPIO-labeled cells. However, drawbacks also exist for these contrast agents. First, the SPIO method requires long incubation times. To overcome this problem, advanced labeling methods such as magnetoelectroporation or magnetosonoporation have been reported (Walczak et al., 2005). In both methods, either a high-voltage electrical pulse or ultrasound was used to incorporate

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SPIO into the cytoplasm for a shorter time with a higher efficiency. Our data, however, demonstrated that electroporation alters the differentiation capability of mouse embryonic stem cells. In addition, the intracellular concentration of SPIO is diluted gradually with cell division or death. Second, at high doses of MnCl2, hESC toxicity occurs at the cellular level and systemic effects on neurologic and cardiovascular functions have been reported. These untoward effects of MnCl2 , however, have been overcome by concurrent calcium supplementation (Bruvold et al., 2005). The most successful optical method for imaging small numbers of transplanted cells in experimental models is bioluminescence. The sensitivity reaches 10−15 to 10−17 mole/liter enabling detection of 100 cells. This modality utilizes an internal biological light source, such as luciferase, which can be detected within the tissues of small animals using sensitive low-light imaging systems. Specific targeting of luciferase transgene expression in restricted cells and tissues of interest has allowed the localization and tracking of cell fate for studying a variety of disease processes. The CCD camera of the BLI is capable of detecting a minimum radiance of 100 photons per second per cm2 per steradian (photons/sec/cm2 /sr) and achieves a minimal image pixel resolution of 50 μm (Wu et al., 2003). High reproducibility (within ±8% SD from mean values) and detection sensitivity of this bioluminescence system for monitoring luciferase reporter gene expression has been demonstrated in vivo (Wu et al., 2001). While optical imaging detects signals from near-cellular level, this technique is limited to small animal imaging due to limited depth penetration of 1 to 2 cm, spatial resolution of 3 to 5 mm, and temporal resolution of

Magnetic Resonance Imaging of hESCs

seconds to minute (Auerbach et al., 1999). Clinical implementation of this technique (bioluminescence imaging) is not feasible.

Critical Parameters Although direct hESC labeling is simple and convenient, incubation time with SPIO and MnCl2 needs to be optimized. Satisfactory labeling of hESC could be achieved in an 8 to 12 hr incubation time with SPIO and in half an hour with MnCl2 without cytotoxicity. Excessive incubation time does not increase SPIO or MnCl2 uptake into cells but it does increases cytotoxicity.

Troubleshooting SPIO labeling is more sensitive than MnCl2 . With the imaging method described above, 50,000 hESC could be visualized with SPIO, while manganese chloride requires ∼106 hESC for direct MR visualization using a 3 Tesla clinical scanner. Care should be taken to make an MRI phantom as homogeneous as possible to remove any potential source of background artifacts such as air bubbles or cracks within the gelatin-based phantom.

Anticipated Results Hypointense, dark signals can be generated from the SPIO-labeled cells on T2*-weighted sequences (Fig. 5A.3.1). A visually distinct contrast can be observed starting at a magnetic field as low as 0.3T. Similarly, remarkable hyperintense, positive signals can be produced from the manganese-labeled hESC using T1weighted sequences (Fig. 5A.3.2). Significant bright signals can be achieved at a magnetic field as low as 1.5T.

Time Consideration

The entire procedure will take ∼72 hr from preparation of cells to MRI scanning of labeled

Figure 5A.3.5 In vivo manganese enhanced MRI of mESC transplanted into the murine right hind limb. The positive signal generated by mESC following intravenous administration of manganese is indicated by a black arrow. This mouse was scanned using a 3T clinical MRI scanner (GE Healthcare System).

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cells. Labeling incubation time can be optimized from 4 to 12 hr with SPIO and from 30 min to 1 hr with MnCl2 . The MnCl2 solution must also be prepared on the day of cell labeling. Excessive incubation time may induce cytotoxicity.

Literature Cited Aoki, I., Takahashi, Y., Chuang, K.H., Silva, A.C., Igarashi, T., Tanaka, C., Childs, R.W., and Korefsky, A.P. 2006. Cell labeling for magnetic resonance imaging with the T1 agent manganese chloride. NMR Biomed. 19:50-59. Auerbach, M.A., Schoder, H., Hoh, C., Gambhir, S.S., Yaghoubi, S., Sayre, J.B., Silverman, D., Phelps, M.E., Schelbert, H.R., and Czernin, J. 1999. Prevalence of myocardial viability as detected by positron emission tomography in patients with ischemic cardiomyopathy. Circulation 99:2921-2926. Bruvold, M., Nordhoy, W., Anthonsen, H.W., Brurok, H., and Jynge, P. 2005. Manganesecalcium interactions with contrast media for cardiac magnetic resonance imaging: A study of manganese chloride supplemented with calcium gluconate in isolated Guinea pig hearts. Invest. Radiol. 40:117-125. Bulte, J.W., Douglas, T., Witwer, B., Zhang, S.C., Strable, E., Lewis, B.K., Zwicke, H., Miller, B., van Geleren, P., Moscovitz, B.M., Duncan, I.D., and Frank, J.A. 2001. Magnetodendrimers allow endosomal magnetic labeling and in vivo tracking of stem cells. Nat. Biotechnol. 19:11411147. Chen, I.Y., Greve, J.M., Gheysens, O., Willmann, J.K., Rodriguez-Porcel, M., Chu, P., Sheikh, A.Y., Faranesh, A.Z., Paulmurugen, R., Yang, P.C., Wu, J.C., and Gambhir, S.S. 2008. Comparison of optical bioluminescence reporter gene and superparamagnetic iron oxide MR contrast agent as cell markers for noninvasive imaging of cardiac cell transplantation. Mol. Imaging Biol. 11:178-187. Frank, J.A., Zywicke, H., Jordan, E.K., Mitchell, J., Lewis, B.K., Miller, B., Bryant, L.H. Jr., and Bulte, J.W. 2002. Magnetic intracellular label-

ing of mammalian cells by combining (FDAapproved) superaramagnetic iron oxide MR contrast agents and commonly used transfection agents. Acad. Radiol. 9:S484-S487. Kostura, L., Kraitchman, D.L., Mackay, E.M., Pittinger, M.F., and Bulte, J.W. 2004. Feridex labeling of mesenchymal stem cells inhibits chondrogenesis but not adipogenesis or osteogenesis. NMR Biomed. 17:513-517. Kraitchman, D.L., Heldman, A.W., Atalar, E., Amado, L.C., Martin, B.J., Pittenger, M.F., Hare, J.M., and Bulte, J.W. 2003. In vivo magnetic resonance imaging of mesenchymal stem cells in myocardial infarction. Circulation 107:2290-2293. Li, Z., Suzuki, Y., Huang, M., Cao, F., Xie, X., Connolly, A.J., Yang, P.C., and Wu, J.C. 2008. Comparison of reporter gene and iron particle labeling for tracking fate of human embryonic stem cells and differentiated endothelial cells in living subjects. Stem Cells 26:864-873. Lin, Y.J. and Koretsky, A.P. 1997. Manganese ion enhances T1-weighted MRI during brain activation: An approach to direct imaging of brain function. Magn. Reson. Med. 38:378-388. Suzuki, Y., Zhang, S., Kundu, P., Yeung, A.C., Robbins, R.C., and Yang, P.C. 2007. In vitro comparison of the biological effects of three transfection methods for magnetically labeling mouse embryonic stem cells with ferumoxides. Magn. Reson. Med. 57:1173-1179. Walczak, P., Kedziorek, D.A., Gilad, A.A., Lin, S., and Bulte, J.W. 2005. Instant MR labeling of stem cells using magnetoelectroporation. Magn. Reson. Med. 54:769-774. Wu, J.C., Sundaresan, G., Iyer, M., and Gambhir, S.S. 2001. Noninvasive optical imaging of firefly luciferase reporter gene expression in skeletal muscles of living mice. Mol. Ther. 4:297306. Wu, J.C., Chen, I.Y., Sundaresan, G., Min, J.J., De, A., Qiao, J.H., Fishbein, M.C., and Gambhir, S.S. 2003. Molecular imaging of cardiac cell transplantation in living animals using optical bioluminescence and positron emission tomography. Circulation 108:1302-1305.

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Lineage Tracing in the Intestinal Epithelium

UNIT 5A.4

Nick Barker1 and Hans Clevers1 1

Hubrecht Institute for Developmental Biology and Stem Cell Research, and University Medical Center Utrecht (UMCU), Utrecht, The Netherlands

ABSTRACT This unit describes the theory and detailed protocols for performing in vivo lineage tracing from Lgr5+ve intestinal stem cells using an Lgr5-EGFP-ires-CreERT2/Rosa26lacZ mouse model. Lineage tracing can be initiated in mice at any age by administering limiting doses of the hormone tamoxifen. This activates the lacZ reporter gene in the Lgr5+ve stem cells, which subsequently transmit this permanent genetic mark to their progeny as they repopulate the epithelium during normal homeostasis. Because the Lgr5+ve cells are long-lived, self-renewing stem cells, they continuously generate lacZ progeny, which contribute to tissue renewal over the entire lifetime of the mouse. The same protocols can be applied to performing in vivo lineage tracing from other Lgr5+ve stem cell populations, including those in the hair-follicle and stomach. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 13:5A.4.1-5A.4.11.  Keywords: Lgr5 r in vivo lineage tracing r stem cell r intestine

INTRODUCTION The biology of the intestine is very well understood, yet the identity of the intestinal stem cells has remained elusive because of a lack of speciÞc markers (Barker et al., 2008). The authors recently identiÞed the Wnt target gene Lgr5 as a speciÞc marker for a restricted population of proliferating cells at the crypt base in both the small intestine and colon (Barker et al., 2007). In the small intestine, these Lgr5+ve cells are wedge-shaped cells called crypt base columnar cells (CBC), which are candidate stem cells found intermingled with the differentiated Paneth cells (Cheng and Leblond, 1974; Bjerknes and Cheng, 1981). To assess the stem cell potential of these Lgr5+ve cells, a mouse model was generated in which an EGFP-ires-CreERT2 expression cassette was inserted immediately downstream of the transcription start site of the endogenous Lgr5 promoter. The Lgr5+ve cells in this mouse consequently express an EGFP tag that allows visualization of them within the intestine using confocal microscopy and also allows us to efÞciently isolate them for in vitro analysis using FACS. The same Lgr5+ve cells also express a tamoxifen-inducible Cre (catalyzes recombination) enzyme that allows for in vivo lineage tracing when the mice are crossed with inducible reporter mice such as Rosa26lacZ (Soriano, 1999; Fig. 5A.4.1). Using this approach, it has been shown that a lacZ reporter gene activated in the Lgr5+ve cells is rapidly inherited by daughter cells, which constantly re-populate the intestinal epithelium as renewal occurs over the lifetime of the animal. In conclusion, this showed that the Lgr5+ve cells are the true stem cells of the intestine. Using a similar approach, it was also shown that Lgr5+ve populations in other tissues such as the hair-follicle (Jaks et al., 2008) and stomach (Barker et al., 2010) are adult stem cell populations responsible for maintaining tissue homeostasis. This unit provides step-by-step instructions for performing this in vivo lineage tracing using the Lgr5-EGFP-ires-CreERT2/Rosa26lacZ mouse model. The preparation and administration of the tamoxifen hormone (see Support Protocol) is described. There are

Current Protocols in Stem Cell Biology 5A.4.1-5A.4.11 Published online May 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a04s13 C 2010 John Wiley & Sons, Inc. Copyright 

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Figure 5A.4.1 Image outlining in vivo lineage tracing using the Lgr5-EGFP-ires-CreERT2/Rosa26-lacZ mice. (A) Intestinal stem cells harboring the Lgr5-EGFP-ires-CreERT2 transgene express both EGFP and the CreERT2 enzyme. In the absence of tamoxifen, the CreERT2 enzyme is sequestered by heat-shock proteins in the cytoplasm. As a result, the lacZ reporter gene remains switched off due to the presence of a transcriptional roadblock. (B) Following induction, tamoxifen is taken up by the Lgr5+ve intestinal stem cells and complexes with the CreERT2 protein in the cytoplasm. This releases the CreERT2 enzyme from its heat-shock chaperones and allows it to enter the nucleus and catalyze the excision of the transcriptional roadblock from the lacZ reporter gene via Cre/loxP-mediated recombination. The lacZ reporter gene is consequently permanently switched on in the Lgr5-EGFP+ve stem cells. (C) These EGFP+ve /lacZ+ve stem cells subsequently divide to generate Lgr5-EGFP−ve progeny, which therefore inherit the genetic lacZ mark. These lacZ+ve progeny rapidly re-populate the intestinal epithelium, allowing their appearance and fate to be tracked over time.

protocols for optimal isolation and Þxation (see Basic Protocol 1) and lacZ staining of the intestines (see Basic Protocol 2). Finally, whole-mount (see Basic Protocol 3) and tissue section (see Basic Protocol 4) protocols for analyzing the lacZ staining are described.

Lineage Tracing in the Intestinal Epithelium

NOTE: All protocols using live animals must Þrst be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow ofÞcially approved procedures for the care and use of laboratory animals.

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ISOLATION AND FIXATION OF THE INTESTINE To visualize lineage tracing in the intestine, it is necessary to dissect, Þx, and lacZ stain the tissue from tamoxifen-treated Lgr5-EGFP-CreERT2/Rosa26lacZ mice.

BASIC PROTOCOL 1

Materials Tamoxifen-treated Lgr5-ires-CreERT2/Rosa26-lacZ mice (see Support Protocol) Gluteraldehyde lacZ Þxative (see recipe) or paraformaldehyde lacZ Þxative (see recipe) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 3-ml syringes and 21-G needles 50-ml centrifuge tube Rolling platform 1. Dissect the intestines from a tamoxifen-treated mouse and place into a petri dish containing 20 ml cold Þxative. 2. Equally divide the freshly isolated intestine into proximal, middle, distal, and colon segments and immediately ßush with 3 ml ice-cold gluteraldehyde Þxative to remove feces using a 3-ml syringe with a 21-G needle attached. An adult mouse small intestine is typically 40-cm long. The proximal segment is deÞned as the 12 to 14 cm immediately adjacent to the stomach. The distal segment is deÞned as the 12 to 14 cm immediately adjacent to the caecum. The intervening 12 to 14 cm deÞnes the middle segment. The colon is typically 7- to 8-cm long and is deÞned as the segment from the caecum to the rectum. The intestine is very susceptible to degradation following death because of its large microbial load. It is therefore crucial to isolate this tissue, ßush thoroughly, and initiate Þxation as soon as possible.

3. Place each intestinal segment into a separate 50-ml centrifuge tube containing 50 ml (∼20-fold excess) of ice-cold gluteraldehyde Þxative. Gluteraldehyde lacZ Þxative generally delivers optimal lacZ stains, but is incompatible with the majority of immunohistochemical procedures. PFA lacZ Þxative can be used in place of gluteraldehyde lacZ Þxative when lacZ/antibody co-stains are required.

4. Fix the intestines by constant mixing on a rolling platform for 2 hr at 4◦ C. Fixation times are critical—over-Þxation can destroy lacZ activity in the tissue.

5. Remove the Þxative and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform. Proceed to Basic Protocol 2.

TAMOXIFEN-INDUCTION OF IN VIVO LINEAGE TRACING In these studies, tamoxifen is used to induce the lineage tracing from the Lgr5+ve intestinal stem cells. The tamoxifen is prepared from a powder and injected into the Lgr5-EGFP/CreERT2/Rosa26lacZ mice.

SUPPORT PROTOCOL

Materials Tamoxifen powder (Sigma, cat. no. T-5648), stored at least 1 year at 4◦ C 100% ethanol Sunßower oil (supermarket variety) Adult Lgr5-iresCreERT2/Rosa26-lacZ mice (6 to 8 weeks old, ∼25 g; Jackson Laboratory) 37◦ C incubator 1-ml syringe and 25-G needle (BD Microlance)

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1. Dissolve the tamoxifen powder at 100 mg/ml in 100% ethanol by extensive vortexing at room temperature. 2. Add sunßower oil to achieve a 10 mg/ml tamoxifen/oil emulsion by extensive vortexing and store 0.4-ml aliquots up to 2 years at −20◦ C. 3. Pre-warm the tamoxifen stock (10 mg/ml) to 37◦ C and thoroughly vortex to ensure a homogenous emulsion. 4. Inject adult Lgr5-ires-CreERT2/Rosa26-lacZ mice intraperitoneally (i.p.) with 100 μl of 10 mg/ml tamoxifen (40 mg/kg) using a 1-ml syringe and 25-G needle. House induced mice under standard conditions. BASIC PROTOCOL 2

β-GALACTOSIDASE (lacZ) STAINING TO VISUALIZE INTESTINAL STEM CELLS The LacZ+ve progeny of the Lgr5+ve stem cells are visualized in the intestine by βgalactosidase staining.

Materials Fixed, freshly isolated intestines from tamoxifen-treated mice (see Basic Protocol 1) Equilibration buffer (see recipe) β-galactosidase (lacZ) substrate (see recipe) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 4% (w/v) paraformaldehyde (PFA; see recipe) Rolling platform 50-ml centrifuge tubes 1. Following the Þnal wash of the intestines (see Basic Protocol 1), remove the CMFPBS from the intestinal sections and add 50 ml equilibration buffer to each tube. Allow the intestines to equilibrate by constant mixing on a rolling platform 30 min at room temperature. 2. Transfer the intestines to a 50-ml centrifuge tube containing 50 ml of freshly-made lacZ substrate and allow the staining reaction to proceed with constant mixing on a rolling platform overnight at room temperature in the dark. X-gal (5-bromo-4-chloro-3-indolyl-β-galactosidase) is light-sensitive and incubation with this substrate and the subsequent post-Þxation in 4% PFA should therefore be performed in the dark.

3. Remove the staining solution and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform in the dark. 4. Remove CMF-PBS and add 50 ml (∼20-fold excess) of 4% PFA to each tube and Þx the intestines by constant mixing on a rolling platform overnight at 4◦ C in the dark. Optimal Þxation is achieved using a minimum 20-fold excess of 4% PFA at 4◦ C.

5. Remove the Þxative and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform. Proceed to Basic Protocol 3. BASIC PROTOCOL 3 Lineage Tracing in the Intestinal Epithelium

WHOLE-MOUNT ANALYSIS OF lacZ STAINING IN THE INTESTINE Whole-mount segments of intestine are examined for the lacZ+ve progeny of the Lgr5+ve stem cells. The tissue is embedded in agarose, sectioned using a vibratome, and analyzed under a stereo microscope.

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Materials Fixed and stained intestinal sections from tamoxifen-treated mice (see Basic Protocol 1) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 4% low-melting-point agarose (Invitrogen, cat. no. 16520-100), pre-warmed to 40◦ C Glue (Bison, http://www.bison.nl, cat. no. Bi2058) Dissection pins Cardboard Petri dishes Stereo microscope (e.g., Olympus SZX9) linked to a digital camera Plastic basemolds (Klinipath, cat. no. 3051-P) Scalpel Vibratome (Microm model HM650V) Vibratome knives (Gillette, cat. no. 10) Starfrost microscope slides Coverslips (Menzel-Gl¨aser) 1. To gain a global view of lacZ staining in the intestine, cut open a 1-cm piece from each intestinal segment along its length and pin it out onto a piece of cardboard with the inner surface (the surface epithelium) facing upwards. 2. Submerge the cardboard in a petri dish containing CMF-PBS and take whole-mount photos of the surface epithelium using a stereo microscope (e.g., Olympus SZX9) linked to a digital camera, with surface illumination (Fig. 5A.4.2). 3. To generate a more detailed overview of the lacZ staining present on local areas of surface epithelium, cut open a ∼1-cm piece of intestine along its length, lay it ßat in a tissue mold, and add 4% low-melting-point agarose until the tissue is completely submerged. 4. Allow the agarose to set (∼20 min), then remove the agarose/tissue block from the mold, and trim it to a perfect square using a scalpel blade. 5. Glue the block to the cutting platform of the vibratome so that the piece of intestine is perpendicular to the knife (i.e., side-on).

A

B

Figure 5A.4.2 Whole-mount analysis of in vivo lineage tracing in the small intestine. (A) Lowpower image showing the presence of multiple lacZ+ve epithelial units throughout the small intestine 600 days post-induction. (B) LacZ+ve epithelial units visible on a 150-μm vibratome section of small intestine 5 days post-induction.

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6. Cut 150- to 200-μm sections (typically containing two to three crypts plus associated surface epithelium) and carefully transfer them to a microscope slide. 7. Place a coverslip over the sections to ensure that they remain ßat and take wholemount pictures as detailed above (Fig. 5A.4.2). BASIC PROTOCOL 4

DETAILED ANALYSIS OF lacZ STAINING ON TISSUE SECTIONS Tissue sections are prepared for closer analysis of lacZ expression.

Materials Intestinal tissue (see Basic Protocol 1) Tissue dehydration solutions: 70%, 80%, 96%, and 100% ethanol n-Butanol (Baker, cat. no. 8017) Liquid parafÞn (60◦ C) De-wax solvent (xylene; Klinipath, cat. no. 4055-9005) Tissue rehydration solutions: 100%, 96%, 90%, 80%, 70%, 60%, 50%, and 25% ethanol 0.1% (w/v) Neutral Red in ddH2 O Pertex mounting medium (Histolab) Tissue cassettes (Klinipath) Metal molds on an embedding station Heated forceps Cold plate (−12◦ C) Microtome 40◦ C water bath Starfrost microscope slides Hot-plate (∼55◦ C) Slide racks (Klinipath) Coverslips (Menzel-Gl¨aser) Digital camera connected to a standard light microscope 1. Transfer the remaining intestinal tissues to a Klinipath tissue cassette and label the front panel clearly using a pencil. 2. Dehydrate the tissues by immersing the cassette in a 20-fold volume of 70% ethanol for 2 hr at 4◦ C. Refresh the 70% ethanol after 1 hr. Repeat this procedure using 96% ethanol and then 100% ethanol. Once tissues are transferred to 70% ethanol, they can be stored for up to 3 months at 4◦ C. The ethanol dehydration series cannot be interrupted after this stage. All dehydration steps using ethanol are performed at 4◦ C to ensure a gradual reduction in hydroxyl (water) bonds within the tissue, thereby reducing tissue damage.

3. Transfer the tissue cassettes to a 20-fold volume of n-butanol and incubate for 2 hr at room temperature. Following dehydration in the ethanol series, n-butanol must be used to remove the last traces of ethanol. Incubation in xylene will result in a loss of lacZ staining in the tissue.

4. Remove the tissue cassettes from the n-butanol and blot them onto tissues to remove any excess solvent. 5. Immerse the cassettes into 60◦ C liquid parafÞn overnight. Lineage Tracing in the Intestinal Epithelium

6. Remove the tissue cassettes from the liquid parafÞn and transfer them into metal molds on an embedding station. Carefully orient the tissues within the parafÞn using

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heated forceps and then transfer the parafÞn blocks to a −12◦ C cold-plate for 30 min to allow them to solidify. 7. Prepare 6-μm thick sections using a microtome and transfer to a clean 40◦ C water bath. Allow the sections to spread out on the water and then ßoat them onto the upper surface of frosted microscope slides. 8. Dry the slides 1 hr on a 55◦ C hot-plate. 9. Transfer the slides into a slide rack and de-wax them two times by immersion in xylene, for 5 min each time. 10. Rehydrate the tissue sections by serial immersion in ethanol as follows:

1 min 1 min 1 min 1 min 1 min 1 min 1 min 1 min 1 min

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100% ethanol (two times) 96% ethanol 90% ethanol 80% ethanol 70% ethanol 60% ethanol 50% ethanol 25% ethanol ddH2 O.

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Figure 5A.4.3 High-resolution examples of short-term and long-term lineage tracing in the small intestine and colon. (A) Isolated Lgr5-lacZ+ve stem cells present at the crypt base in the small intestine 1 day post-induction (black arrow). (B) Epithelial units in the small intestine partially populated by lacZ+ve progeny 5 days post-induction. LacZ+ve Paneth cells are typically not observed at these early time-points (red arrows). (C) Epithelial units in the small intestine entirely populated by lacZ+ve progeny 128 days post-induction. (D) Isolated Lgr5-lacZ+ve stem cells present at the colon crypt base 1 day post-induction (black arrow). (E) Epithelial units in the colon partially populated by lacZ+ve progeny 5 days post-induction. (F) Epithelial units in the colon entirely populated by lacZ+ve progeny 128 days post-induction.

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11. Counterstain by immersion in 0.1% Neutral Red solution for 1 min. Neutral Red is a counterstain that colors both the nucleus and cytoplasm red, generating an optimal contrast against the blue lacZ stain.

12. Quickly rinse the slides in 100% ethanol, three times for 30 sec each time, to remove excess Neutral Red counterstain and then transfer to xylene, incubate two times for 2 min each time. 13. Place a coverslip over the sections and seal it in place using Pertex mounting medium. 14. Photograph the sections using a digital camera connected to a standard light microscope (Fig. 5A.4.3).

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

β-galactosidase (lacZ) substrate 5 mM K3 Fe(CN)6 (see recipe) 5 mM K4 Fe(CN)6 ·3H2 O (see recipe) 2 mM MgCl2 (see recipe) 0.02% (v/v) NP40 (see recipe) 0.1% (v/v) sodium deoxycholate (see recipe) 1 mg/ml X-gal in CMF-PBS (see recipe) Prepare fresh and keep in the dark at room temperature Ethylene glycol tetraacetic acid Prepare 500 mM ethylene glycol tetraacetic acid (EGTA; Sigma) in ddH2 O. Adjust the pH to 7.2 using NaOH. Store indeÞnitely at room temperature.

Equilibration buffer 2 mM MgCl2 (see recipe) 0.02% (v/v) NP40 (see recipe) 0.01% (w/v) sodium deoxycholate in CMF-PBS (see recipe) Store indeÞnitely at room temperature Gluteraldehyde lacZ Þxative 1% (v/v) formaldehyde 0.2% (v/v) gluteraldehyde 0.02% (v/v) NP40 in CMF-PBS (see recipe) Prepare fresh and keep on ice Magnesium chloride Prepare a 1 M magnesium chloride (MgCl2 ) stock in CMF-PBS. Store indeÞnitely at room temperature.

NP40 Prepare a 10% (v/v) NP40 stock in CMF-PBS. Store indeÞnitely at room temperature.

Paraformaldehyde, 4% (w/v) Lineage Tracing in the Intestinal Epithelium

Dissolve 40 g paraformaldehyde (PFA; Sigma, cat. no. P6148) per liter of CMFPBS and heat to 60◦ C with constant stirring. Store up to 2 weeks (short-term) at 4◦ C or up to 6 months (long-term) at –20◦ C.

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Paraformaldehyde lacZ Þxative 4% (w/v) paraformaldehyde (PFA; see recipe) 15 mM EGTA, pH 7.2 (see recipe) 2 mM MgCl2 in CMF-PBS (see recipe) Prepare fresh and keep on ice Potassium hexacyanoferrate II trihydrate (K4 Fe(CN)6 ·3H2 O) Prepare a 200 mM potassium hexacyanoferrate II trihydrate (K4 Fe(CN)6 ·3H2 O; Sigma, cat. no. P3289) stock in CMF-PBS. Store up to 2 weeks at 4◦ C.

Potassium hexacyanoferrate III (K3 Fe(CN)6 ) Prepare a 200 mM potassium hexacyanoferrate III (K3 Fe(CN)6 ; Sigma, cat. no. P8131) stock in CMF-PBS. Store up to 2 weeks at 4◦ C.

Sodium deoxycholate Prepare a 10% (w/v) sodium deoxycholate stock in ddH2 O. Store indeÞnitely at room temperature.

X-gal Prepare a 50 mg/ml X-gal (5-bromo-4-chloro-3-indolyl-β-galactosidase; Invitrogen, cat. no. 15520-018) stock in dimethylformamide (Sigma ACS grade 3, cat. no. 19937), dispense into 5-ml aliquots, and store up to 6 months at −20◦ C.

COMMENTARY Background Information The inner lining of the small intestine is arranged into multiple functional units of columnar epithelium comprising Þnger-like villi that protrude into the gut lumen, surrounded by ßask-shaped invaginations called crypts of Leiberkuhn (Stappenbeck et al., 2003; Sancho et al., 2004). These villi serve to maximize the surface area available for efÞcient absorption of digested food and water exiting the stomach. In the large intestine (colon), a ßat surface epithelium replaces these villi, reßecting its primary role in compacting undigested food remnants into stool/feces rather than absorption. The intestinal epithelium is probably the most rapidly renewing tissue in adults, undergoing a complete cycle of renewal every 5 days. This self-renewal is driven by a small population of self-renewing, multipotent stem cells located at the base of crypts (Bjerknes and Cheng, 1999). These stem cells generate a transient population of rapidly proliferating cells (the transit amplifying cells) that divide two to three times as they migrate upwards before differentiating into the major cell types present on the surface epithelium as they exit the crypts. These differentiated cells (comprising absorptive enterocytes, mucus-secreting goblet cells, and much rarer

hormone-secreting enteroendocrine cells) perform their essential functions as they continue migrating along the surface epithelium until Þnally dying by programmed cell death after 5 days. A fourth differentiated cell type in the small intestine called the Paneth cell escapes this upward migration and instead differentiates as it moves to the very base of the crypt. This cell lives up to 8 weeks and secretes antimicrobial lysozyme and cryptdins as part of the gut immune system. Lysozyme-secreting Paneth cells are not present in the colon, although cells with a similar function are thought to be present. In vivo lineage tracing in the Lgr5-iresCreERT2/Rosa26-lacZ mouse model centers on the inducible activation of a stablyintegrated lacZ reporter gene in the Lgr5+ve stem cell populations. Daughter cells derived from the lacZ+ve Lgr5 populations inherit this genetic lacZ mark, allowing their appearance and fate to be tracked in the corresponding tissue over time. This approach has been successfully used to demonstrate the stem cell function of Lgr5+ve cells in adult small intestine, colon, hair-follicle, and pyloric stomach. In the absence of tamoxifen induction, the Cre-ERT2 enzyme present exclusively in the Lgr5+ve stem cells is efÞciently sequestered by heat-shock proteins in the cytoplasm. The

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Rosa26lacZ reporter gene in the nucleus consequently remains switched off by virtue of the presence of a transcriptional roadblock sequence ßanked by loxP sites (Fig. 5A.4.1). Intraperitoneal injection of limiting doses of tamoxifen releases the Cre-ERT2 enzyme from its heat-shock protein chaperones and allows it to accumulate in the nucleus of isolated Lgr5+ve cells. Here, it rapidly mediates excision of the transcriptional roadblock by catalyzing recombination across the ßanking LoxP sites. The lacZ reporter gene is thereby activated in the Lgr5+ve cells and subsequently transmitted to their progeny as they repopulate the tissue during normal homeostasis (Barker and Clevers, 2007; Fig. 5A.4.1). Because the Lgr5+ve cells are long-lived, self-renewing stem cells, they continuously generate lacZ progeny, which contribute to tissue renewal over the entire lifetime of the mouse. LacZ reporter gene activity is visualized within the tissue using a 5-bromo-4-chloro3-indolyl-β-galactosidase substrate, which is catalyzed to a blue product by the lacZ enzyme.

Critical Parameters and Troubleshooting

Lineage Tracing in the Intestinal Epithelium

In the Lgr5-EGFP-ires-CreERT2 mouse intestine, clusters of crypts silence the knock-in allele in a region-dependent fashion. The percentage of EGFP-ires-CreERT2+ve crypts consequently decreases from the proximal small intestine (∼70%) to the distal small intestine (∼30%). Such variegated expression of transgenes is commonly observed in the intestine. Importantly, however, no variegated expression is observed in other Lgr5+ve tissues, including the skin and stomach (nor is it observed in the intestine of an independent Lgr5-lacZ knockin model that the authors have used during the course of our studies). The Lgr5-EGFP-ires-CreERT2 mice are only viable as heterozygotes (i.e., one wildtype allele must always be present). In vivo lineage tracing is therefore always performed with Lgr5-EGFP-ires-CreERT2het /Rosa26lacZhet/hom mice. In principle, in vivo lineage tracing can be performed using Lgr5-EGFP-ires-CreERT2 mice in combination with any inducible reporter mouse strain that is activated using standard Cre/loxP technology. The dose of tamoxifen used to induce adult Lgr5-EGFP-ires-CreERT2/Rosa26-lacZ mice (estimated at 25 g total weight) is selected to achieve stochastic activation of the lacZ reporter gene within the Lgr5+ve population

of the intestine (to demonstrate that a single Lgr5+ve stem cell is responsible for driving the renewal of all cell types present on the crypt/villus epithelium). Higher doses of tamoxifen may be used to increase the frequency of lineage tracing if desired.

Anticipated Results

LacZ+ve Lgr5 stem cells at the crypt base should Þrst be observed 12 to 16 hr after induction. LacZ+ve Lgr5-derived transit amplifying cells will subsequently become visible in the crypts within 2 to 3 days. Expect to observe lacZ+ve progeny throughout the crypts and the associated surface epithelium 7 to 10 days postinduction. Typically, lacZ+ve enterocytes (alkaline phosphatase-positive) and goblet cells (PAS-positive) are present at this time-point because these exhibit the highest rate of turnover. Paneth cells (lysozyme-positive) have a much lower turnover rate and lacZ+ve examples are typically observed in the smallintestine only 3 to 4 weeks post-induction. At these early time-points, the surface epithelium in the small intestine contains both lacZ+ve and lacZ−ve progeny, creating a mosaic pattern (Fig. 5A.4.3B). This occurs because the surface epithelium is being supplied with cells from multiple crypts containing both lacZ+ve and lacZ−ve stem cells. After 60 to 80 days, the entire surface epithelium of tracing units is typically comprised of lacZ+ve cells, reßecting the fact that the lacZ+ve stem cell population has become dominant (with respect to the lacZ−ve stem cells) in the crypts supplying this intestinal unit. Given that the Lgr5+ve cells are long-lived, self-renewing stem cells, the frequency of lacZ+ve epithelial units present will remain constant even after the intestinal epithelium has undergone multiple rounds of complete renewal. Thus, expect to see multiple tracing units even 24 months after induction.

Time Considerations Following isolation of the intestines, the entire lacZ staining/Þxation/embedding procedure should take ∼4 days. Once embedded in parafÞn the tissues can be stored long-term at room temperature without loss of lacZ stain.

Literature Cited Barker, N. and Clevers, H. 2007. Tracking down the stem cells of the intestine: Strategies to identify adult stem cells. Gastroenterology 133:1755760.

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Barker, N., van Es, J.H., Kuipers, J., Kujala, P., van den Born, M., Cozijnsen, M., Haegebarth, A., Korving, J., Begthel, H., Peters, P.J., and Clevers, H. 2007. IdentiÞcation of stem cells in small intestine and colon by marker gene Lgr5. Nature 449:1003-1007. Barker, N., van de Wetering, M., and Clevers, H. 2008. The intestinal stem cell. Genes Dev. 22:1856-1864. Barker, N., Huch, M., Kujala, P., van de Wetering, M., Snippert, H.J., van Es, J.H., Sato, T., Stange, D.E., Begthel, H., van den Born, M., Danenberg, E., van den Brink, S., Korving, J., Abo, A., Peters, P.J., Wright, N., Poulsom, R., and Clevers, H. 2010. Lgr5(+ve) stem cells drive self-renewal in the stomach and build longlived gastric units in vitro. Cell Stem Cell 6:2536. Bjerknes, M. and Cheng, H. 1981. The stem cell zone of the small intestinal epithelium III. Evidence from columnar, enteroendocrine, and mucosal cells in the adult mouse. Am. J. Anat. 160:77-91.

Cheng, H. and Leblond, C.P. 1974. Origin, differentiation, and renewal of the four epithelial cell types in the mouse small intestine. V. Unitarian theory of the origin of the four epithelial cell types. Am. J. Anat. 141:537-561. Jaks, V., Barker, N., Kasper, M., van Es, J.H., Snippert, H.J., Clevers, H., and Toftgard, R. 2008. Lgr5 marks cycling, yet long-lived, hair follicle stem cells. Nat. Genet. 40:12911299. Sancho, E., Batlle, E., and Clevers, H. 2004. Signaling pathways in intestinal development and cancer. Annu. Rev. Cell Dev. Biol. 20:695723. Soriano, P. 1999. Generalized lac-Z expression with the ROSA26 Cre expression strain. Nat. Genet. 21:70-71. Stappenbeck, T.S., Mills, J.C., and Gordon, J.I. 2003. Molecular features of adult mouse small intestinal epithelial progenitors. Proc. Natl. Acad. Sci. U.S.A. 100:1004-1009.

Bjerknes, M. and Cheng, H. 1999. Clonal analysis of intestinal epithelial progenitors. Gastroenterology 116:7-14.

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Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes in Mammalian Neocortical Neural Progenitors

UNIT 5A.5

Janice H. Imai,1,2 Xiaoqun Wang,1 and Song-Hai Shi1,2 1

Developmental Biology Program, Memorial Sloan-Kettering Cancer Center, New York, New York 2 BCMB Allied Program, Weill Cornell Medical College, New York, New York

ABSTRACT The importance of the centrosome in regulating basic cellular processes and cell fate decisions has become increasingly evident from recent studies tracing the etiology of developmental disorders to mutations in genes encoding centrosomal proteins. This unit details a protocol for a ßuorescence-based pulse labeling of centrioles of neural progenitor cells in the developing neocortex of mice. In utero electroporation of KaedeCentrin1 followed by in utero or ex vivo photoconversion allows a direct monitoring of the inheritance of centrosomes containing centrioles of different ages in dividing neocortical neural progenitors (i.e., radial glial cells). This is achieved by combining the irreversible photoconversion capacity of the Kaede protein from green to red ßuorescence with the faithful duplication of the centrosome during each cell cycle. After two mitotic divisions following photoconversion, mother centrosomes containing the original labeled centriole appear in both red and green ßuorescence, and can be distinguished from daughter centrosomes which appear in green ßuorescence only. This facilitates the study of the inheritance and behavior of the mother and daughter centrosomes in asymmetric cell divisions in the developing mammalian neocortex. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 15:5A.5.1-5A.5.12.  Keywords: centrosome r Kaede-Centrin1 r mother and daughter centrosomes r photoconversion r neocortex r radial glia progenitor cell r in utero electroporation

INTRODUCTION This unit details a protocol for labeling the centrioles of neural progenitor cells (i.e., radial glia) in the developing neocortex of mice during the peak period of cortical neurogenesis (gestational days 13.5 through 17.5, i.e., E13.5 through 17.5), so as to study centrosome segregation in the context of neurogenesis (Wang et al., 2009). Centrioles/centrosomes of radial glia are initially labeled with green ßuorescence by in utero electroporation of a plasmid bearing the photoconvertible ßuorescent protein (Kaede) fused to the centriolar protein Centrin1 (Kaede-Centrin1) into the developing neocortex of embryos at E13.5. The plasmid is taken up by the radial glia within the ventricular zone. Approximately 24 hr later, each electroporated embryo is exposed to a brief pulse of violet light while in the uterus to convert the Kaede-Centrin1 protein from green to red ßuorescence. The uterus is placed back into the mouse, and the embryos are allowed to continue development. About 48 hr later, the brains of the embryos are recovered and two distinct populations of centrosomes can be observed: one has both green and red ßuorescence, representing the more mature mother centrosomes, and the other has green ßuorescence only, representing the less mature daughter centrosomes. We have shown that mother centrosomes are preferentially inherited by the renewing radial glia remaining in the ventricular zone, while daughter centrosomes are mostly inherited by the differentiating progeny that migrate away from the ventricular zone and occupy more Current Protocols in Stem Cell Biology 5A.5.1-5A.5.12 Published online October 2010 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc05a05s15 C 2010 John Wiley & Sons, Inc. Copyright 

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dorsal layers of the neocortex, including the cortical plate (Wang et al., 2009). These results suggest that centrosome inheritance is tightly regulated and coordinated with cell fate decisions during asymmetric division of neural progenitors in the developing neocortex. This unit Þrst describes the well established method of in utero electroporation of plasmid DNA into radial glia in the developing neocortex (Basic Protocol 1). Next, a method for photoconverting Kaede-Centrin1 in vivo is described (Basic Protocol 2), followed by a procedure for preserving and visualizing the centrosomes with different ßuorescence spectra. Finally, an alternative procedure for photoconverting Kaede-Centrin1 in organotypic neocortical slices for time-lapse imaging studies of centrosome regulation during neurogenesis is presented (Alternate Protocol). NOTE: This protocol was developed in mice, and therefore some parameters must be determined empirically when applied to other species. NOTE: All solutions and equipment contacting the embryos should be sterile. NOTE: All protocols using live animals must Þrst be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow ofÞcially approved procedures for care and use of laboratory animals.

BASIC PROTOCOL 1

IN UTERO ELECTROPORATION OF THE KAEDE-CENTRIN1 PLASMID This protocol has been previously described (e.g., Tabata and Nakajima, 2008) and extensively employed in studying the function of gene(s) of interest in mammalian neocortical development.

Materials Timed-pregnant female mouse, E13.5 Isoßuorane Ethanol and iodine wipes Phosphate-buffered saline, sterile, 37◦ C Plasmid DNA (Kaede-Centrin1, 3.0 μg/μl; contact S.-H. Shi, [email protected]) mixed with Fast Green dye (Fisher Biotech) (1% w/v in PBS, 1 μl dye solution per 10 μl DNA solution); the use of endotoxin-free plasmid DNA (e.g., prepared using Qiagen Endotoxin-free Maxiprep kit) is recommended Antibiotic-PBS solution: penicillin (100 IU/ml)/streptomycin (100 mg/ml) in PBS, warmed to 37◦ C Antibiotic/analgesic solution (Duane Reade Triple Antibiotic Ointment plus Pain Relief; contains Bacitracin, zinc, neomycin sulfate, polymyxin B sulfate, and the analgesic pramoxine)

Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

Isoßurane induction chamber (VetEquip, e.g., cat. no. 901807) Isoßuorane dispenser (VetEquip) and nose cone Heating pad Disposable underpads Hair clipper Surgical instruments 10-ml sterile syringe Sterile gauze Sterile spatula Glass capillary injection needles (tip diameter ∼100 μm; beveled; see recipe) Electroporation system (BTX, ECM830, Harvard Apparatus)

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Silk or nylon sutures Wound clips (7-mm; CellPoint ScientiÞc) Cotton-tipped applicators Surgically prepare the embryos 1. Anesthetize a timed-pregnant female mouse at 13.5 days of gestation (i.e., embryonic day 13.5 or E13.5) with isoßuorane in an induction chamber at a ßow rate of 1 to 4 liters/min (or according to the dispenser manufacturer’s recommendations). A single anesthetization procedure is carried out: Þrst, the animal is placed in the induction chamber until it is breathing but no longer moving; the air ßow valve from the isoßuorane dispenser to the chamber is closed while the air ßow valve from the dispenser to the nose cone is opened, and the animal is quickly transferred from the chamber to the nose cone (on the heating pad) while it is still unconscious.

2. Transfer the mouse from the isoßuorane induction chamber to the surgery table and onto a heating pad (37◦ to 40◦ C) covered with a clean disposable underpad. Place the mouse’s head in a nose cone connected to the isoßuorane output tube with the ßow rate of the isoßuorane set at 0.5 liter/min (or according to the manufacturer’s recommendations). 3. When the mouse is unresponsive to toe pinches, remove the abdominal fur with a hair clipper. Clean the shaven skin with iodine and alcohol wipes.

A

E E

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C C

F

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G G

Figure 5A.5.1 Procedure for in utero electroporation. (A) Timed-pregnant female mouse at gestational/embryonic day 13.5 (E13.5) under isoßuorane anesthesia with abdomen shaven and cleaned. The skin and underlying muscle have been cut open. (B) Mouse covered with a square of sterile gauze with a central hole cut out to expose only the opened abdominal cavity. The uterine horn containing embryos on the right side of the mouse has been gently lifted out of the cavity and placed on the gauze. (C) Positioning the embryo for injection by using a spatula to gently roll the embryo around within the yolk sac. (D) Glass capillary micropipet Þlled with plasmid DNA-dye mixture penetrating the lateral ventricle of the embryo through the uterine wall and yolk sac. (E) Positioning the electrodes for pulse delivery with the positive electrode covering the injected area and the negative electrode contacting the embryo at a diametrically opposed location. (F) Suturing the abdominal muscle after placing the uterine horn back into the abdominal cavity. (G) Applying wound clips to the skin.

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4. Make a vertical incision (∼2.5 cm) in the abdominal skin. To facilitate subsequent suturing of the muscle, separate the skin from the underlying muscle by cutting away the connective tissue. 5. Make a slightly smaller incision in the muscle (Fig. 5A.5.1A). From this point forward, the exposed abdominal tissue as well as the embryos should be continually moistened with warm (37◦ C) PBS dispensed dropwise from a 10-ml syringe.

6. Place a sterile gauze with a central hole cut out upon the abdomen such that only the incision area is exposed. 7. Pull the uterine horns gently out of the abdominal cavity and place on the gauze (Fig. 5A.5.1B).

Inject and electroporate the plasmid 8. Locate the lateral ventricles of the embryonic brain. At E13.5, the lateral ventricles of the developing brain occupy a large portion of each brain hemisphere and can be discerned through the uterine wall and yolk sac as a slightly darker, crescent-shaped area.

9. Gently rotate the embryo within its yolk sac with the aid of a spatula so as to position the head at an optimal angle for injection (Fig. 5A.5.1C). Take care not to squeeze the embryos. 10. Inject ∼1 μl plasmid DNA–dye mixture into the lateral ventricle of each embryo using a beveled glass micropipet (Fig. 5A.5.1D). As the DNA-dye mixture Þlls the ventricle, the ventricle will become more visible as a green crescent shape. In order to efÞciently perform the photoconversion (Basic Protocol 2) the following day and minimize surgery time, it is advisable to make the injection on the same side for each embryo. Only one ventricle per embryo is injected; there is no control per se.

11. After injection, pulse the embryo with a train of Þve 40- to 50-mV pulses (duration: 50 msec; interval: 950 msec) by covering the injected area with the positive electrode while the negative electrode maintains contact with the embryo on the diametrically opposite side of the head or body (Fig. 5A.5.1E). Radial glia line the ventricle in the area known as the ventricular zone; therefore, these take up the plasmid DNA when the voltage pulses are applied. As development proceeds, radial glia divide asymmetrically to produce radial glia that remain in the ventricular zone, as well as more fate-restricted daughter cells that migrate radially away from the ventricular zone to occupy more dorsal layers of the cortex. Care should be taken to avoid contact between the electrodes and the placenta during pulse application.

12. Repeat for each embryo. Generally, all embryos in a litter are injected.

Complete the surgery 13. Place the uterine horns back into the abdominal cavity and bathe in antibiotic-PBS solution. Suture and clip the wound (Fig. 5A.5.1F,G). Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

14. Apply analgesic ointment with a cotton-tipped applicator to the wound area. Place the mouse separately in a clean cage and closely monitor for respiratory distress until it is alert and ambulatory.

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IN UTERO PHOTOCONVERSION OF THE KAEDE REPORTER Before undertaking this protocol, the minimum time required for complete photoconversion of Kaede-Centrin1 from green to red ßuorescence should be tested. For this calibration procedure, embryos are injected at E13.5, and the following day (E14.5) the embryos are exposed to violet light (see below); ∼3 to 5 min is usually sufÞcient. The light beam will penetrate the uterine wall, the yolk sac, and the outer tissues of the embryo’s head to the deepest layer of the developing neocortex, i.e., the ventricular zone where the Kaede-Centrin1-expressing radial glia are located. Embryos can be sacriÞced immediately following the photoconversion in these pilot experiments to examine the efÞcacy of the photoconversion.

BASIC PROTOCOL 2

Materials Pregnant mouse with electroporated embryos at E13.5 (Basic Protocol 1) Source of violet light (e.g., a ßuorescence dissection microscope with a mercury lamp and a 4 ,6-diamidino-2-phenylindole [DAPI] Þlter) Wound clip remover Additional reagents and equipment for anesthesia of the mouse and surgically removing and replacing embryos (Basic Protocol 1) 1. Approximately 24 hr after in utero electroporation, anesthetize the mouse with isoßuorane (see Basic Protocol 1) and reopen the wound by removing wound clips and sutures. 2. Expose the uterine horns and hold each embryo under the beam of a violet light (350 to 400 nm), which is focused on the injected area, for 3 to 5 min (Fig. 5A.5.2). 3. After all embryos are treated, place the uterine horns back into the abdominal cavity and resuture and clip the wound. 4. Apply appropriate analgesia and closely monitor the mouse until it has recovered, as described in Basic Protocol 1.

violet light beam

Figure 5A.5.2 Setup for in utero photoconversion. The embryo is held under a beam of violet light to effect photoconversion. The beam of light covers the injected area.

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ALTERNATE PROTOCOL

EX VIVO PHOTOCONVERSION IN BRAIN SLICES AND TIME-LAPSE IMAGING The same principles used for in vivo photoconversion can be applied to organotypic brain slice cultures, when the desired goal is a time-lapse imaging study of mother and daughter centrosome behavior over the course of neocortical neurogenesis. As for Basic Protocol 2, the minimum time required for complete photoconversion should be determined for slices in pilot experiments. Usually a much shorter time (<1 min) is required to achieve near-complete photoconversion in slice cultures. The protocol for preparing organotypic neocortical slice cultures has been previously described (Daza et al., 2007; Elias and Kriegstein, 2007).

Additional Materials (also see Basic Protocols 1 and 2) Pregnant mouse with electroporated embryos at E13.5 (Basic Protocol 1) 4% (w/v) agarose in artiÞcial cerebrospinal ßuid (ASCF; see recipe) Brain slice culture medium (see recipe) Vibratome (Leica Microsystems) Slice culture insert (Millicell, Millipore) Glass-bottom petri dish (MatTek Corporation) Inverted microscope (e.g., Axiovert 200, Zeiss) with a mercury lamp and a DAPI Þlter HumidiÞed incubator (37◦ C and 5% CO2 ) 1. Perform in utero electroporation of Kaede-Centrin1 at E13 (Basic Protocol 1). 2. One day later, dissect the brain from each live embryo (Basic Protocol 1) and embed in 4% agarose in artiÞcial cerebrospinal ßuid (ACSF). Dissolve 4 g of low-melt agarose (Fisher, cat. no. BP1360-100) in 100 ml of ACSF in an Erlenmeyer ßask in a microwave oven. Once the agarose is dissolved, the ßask is placed in a 42◦ C water bath until the temperature of the agarose reaches ∼42◦ C (∼30 min). The brain is rapidly dissected from each live embryo and placed in ice-cold ACSF for 5 min, then quickly transferred to the agarose, which has been poured into a 6-mm petri dish. The brain is gently positioned in the agarose such that coronal sections can be made easily (i.e., dorsal side up and ventral side down). When the agarose has solidiÞed, the brain is cut out as a cube and attached with a highly adhesive glue to the specimen plate of the Vibratome. IMPORTANT NOTE: The specimen chamber and glue used for cutting live brains should be separate from those used for Þxed brains.

3. Cut coronal sections of the neocortex (∼300 to 400 μm) using a Vibratome. 4. Place the sections onto membrane inserts (Millicell, Millipore) in a 35-cm glassbottom Petri dish with 750 μl of brain slice culture medium. Maintain the cultures in a humidiÞed incubator (37◦ C and 5% CO2 ) for days. 5. For photoconversion, transfer the dish containing the organotypic culture slices to an inverted microscope (e.g., Axiovert 200, Zeiss) with a mercury lamp and a DAPI Þlter. Expose the slices to a brief (i.e., hundreds of milliseconds to a few seconds) pulse of epißuorescent illumination using a DAPI Þlter. 6. Return the slices to the incubator. 7. Take images of labeled centrosomes at the desired time point. Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

The clear segregation of mother and daughter centrosomes can be witnessed after 1.5 to 2 divisions of the radial glia, i.e., ∼48 hr after photoconversion.

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TISSUE PRESERVATION, IMAGING, AND ANALYSIS Images of mother and daughter centrosomes are obtained after photoconversion of the Kaede-Centrin1. Labeled brains are dissected from perfused embryos, post-Þxed, sectioned, and imaged by confocal laser scanning microscopy.

SUPPORT PROTOCOL

Materials 4% Avertin (2,2,2-tribromoethanol; Sigma) in PBS 4% paraformaldehyde (PFA), freshly prepared (cold) Pregnant mouse with electroporated embryos that have been subjected to photoconversion (Basic Protocol 2) Phosphate-buffered saline (PBS), pH 7.4 (cold) 4% (w/v) paraformaldehyde in PBS, freshly prepared 3% to 4% (w/v) agarose in PBS PBS with 0.03% (w/v) sodium azide 1-ml syringe with 30-G, 1-in. needle for Avertin injection Dissection dish (60-mm Pyrex dish lined with 7 to 8 mm of Sylgard 184 Silicone Elastomer) Insect pins (Fine ScientiÞc Tools) Dissection microscope Dissection tools for removing brain (Þne forceps, scissors, spatula) Two 10-ml syringes connected by a stopcock, with 30-G, 0.5-in needle attached to tubing (Fig. 5A.5.3A) Vibratome (Leica Microsystems) 48-well culture plate Microscope equipped for confocal laser scanning microscopy (Olympus FV1000) Remove embryos from uterus 1. At 48 hr after photoconversion (Basic Protocol 2), sacriÞce the mouse by intraperitoneal injection of a lethal dose of Avertin (∼0.8 ml). 2. When the animal is unresponsive to toe pinches but still breathing, cut open the abdominal cavity and expose the uterine horns. 3. Remove each embryo from its yolk sac (Fig. 5A.5.3B) and pin to a dissection dish (Fig. 5A.5.3C) under a dissecting microscope.

Fix embryos by perfusion 4. To transcardially perfuse the embryo, insert the perfusion needle into the left ventricle of the beating heart, and make a small incision to the right atrium to permit the outßow of blood (Fig. 5A.5.3D). 5. Slowly dispense cold PBS slowly from the syringe (3 to 4 ml is usually sufÞcient for E16.5 embryos) until the outßow of blood from the right atrium has ceased and the embryo is white (Fig. 5A.5.3E). 6. Adjust the stopcock to the open position for freshly prepared 4% (w/v) paraformaldehyde in PBS, and infuse a volume of that solution similar to the amount of PBS used in step 5 through the heart.

Dissect out and section brain 7. Dissect out the brain (Fig. 5A.5.3F,G) and post-Þx overnight at 4◦ C with 4% PFA in PBS. 8. The following day, rinse the brain with PBS, and embed in 3% to 4% agarose in PBS.

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A A

B B

needle

F F

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C C

G G RA LA RV

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Figure 5A.5.3 Set-up and procedure for embryo dissection, transcardial perfusion, and brain dissection. (A) Syringes containing cold phosphate-buffered saline (PBS) and 4% paraformaldehyde (PFA) connected by a two-way stopcock with attached tubing and needle for transcardial perfusion. (B) Dissection of embryo (gestational day 16.5) from uterus of anesthetized mouse (C to E, ventral view). (C) Dissected embryo pinned facing upward to a dissection dish Þlled with cold PBS. (D) Close-up view of exposed but intact heart with arrows to indicate the ßow of perfusion solutions into the left ventricle (LV) and out of the right atrium (RA). (E) Embryo after perfusion; heart chambers are outlined by a broken white line. Note the white color of the head when perfusion is complete. LA, left atrium; RV, right ventricle. (F and G, dorsal view). (F) Perfused embryo pinned facing downward, with cranial skin and bone being removed with Þne forceps. (G) Exposed brain being removed with spatula. The left (LH) and right (RH) hemispheres of the brain are outlined by a broken white line.

Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

Dissolve 4 g of low-melt agarose (Fisher, cat. no. BP1360-100) in 100 ml of PBS in an Erlenmeyer ßask by heating in a microwave oven. Once the agarose is dissolved, the ßask is placed in a 42◦ C water bath until the temperature of the agarose reaches ∼42◦ C (∼30 min). The brain is placed in the agarose, which has been poured into a 6-mm petri dish. The brain is gently positioned in the agarose in such a way as to facilitate coronal sectioning (i.e., dorsal side up and ventral side down). When the agarose has solidiÞed, the brain is cut out as a cube and glued to the specimen plate of the Vibratome.

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9. Section the brain at 50 to 80 μm in the coronal plane using a Vibratome. 10. Collect ßoating sections in the wells of a 48-well plate (each well containing 1 ml PBS with 0.03% w/v sodium azide).

Image centrosomes 11. Image centrosomes by confocal laser scanning microscopy using a 40× or 60× objective. Alternatively, sections can be mounted onto glass slides and covered with mounting medium and coverslip for imaging.

REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

ArtiÞcial cerebrospinal ßuid (ACSF; pH 7.4, 310 mOsm/liter) 125 mM NaCl 5 mM KCl 1.25 mM NaH2 PO4 1 mM MgSO4 2 mM CaCl2 25 mM NaHCO3 20 mM glucose Prepare fresh on day of use Brain slice culture medium 50% (v/v) Eagle’s Basal Medium (Invitrogen) 25% (v/v) Hanks’ balanced salt solution 5% (v/v) fetal bovine serum (FBS) 1% (v/v) 100× N2 supplement (Invitrogen) 1% (v/v) 100× penicillin/streptomycin (Invitrogen) 1% (v/v) 100× glutamine (Invitrogen) 0.66% (w/v) D-(+)-glucose (Sigma) Store up to 2 weeks at 4◦ C Glass capillary injection needles To make injection needles, starting with the micropipettes (PCR Micropipets 1 to 10 μl; Drummond, cat. no. 5-000-1001-X10), we use a Flaming/Brown Micropipette Puller (Sutter Instruments, Model P-97) at the following settings: P = 350; Heat = 630; Pull = 24; Vel. = 40; Time = 40. We break the tip of the needle with Þne forceps so that the tip diameter is roughly 60 μm, then polish the tip to a sharp point using a K.T. Brown Type Micropipette Beveler (Sutter Instruments, Model BV-10). Detailed instructions on making pipettes are available in Sutter Instruments’ “P-1000 & P-97 Pipette Cookbook, 2010 Rev. F.”

COMMENTARY Background Information The discovery and characterization of many novel centrosomal proteins in recent years has shed light on the varied and essential functions of the centrosome in controlling cell behaviors and cell fate decisions during development (Schatten and Sun, 2010) and in

disease (Nigg and Raff, 2009). These functions may be especially important in a complex tissue such as the mammalian neocortex, a multilayer structure whose organization depends upon a tightly regulated balance between progenitor cell self-renewal and differentiation.

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The neocortex begins as a layer of neuroepithelial cells surrounding the lateral ventricles at around E9.5 in mice (Molyneaux et al., 2007). Neuroepithelial cells divide to give rise to radial glia, a major population of neural progenitors in the developing neocortex; these cells undergo asymmetric division to self-renew and also to produce more fate-

Kaede-Centrin1 red

Kaede-Centrin1 green

restricted cell types, including neurons (G¨otz and Huttner, 2005). The precise mechanisms that direct the asymmetric division of radial glia are not entirely clear; however, the differential inheritance of mother and daughter centrosomes by radial glia and their more differentiated daughter cells has recently been observed in

␥-tubulin

overlay

Figure 5A.5.4 Kaede-Centrin1 labeling of centrosome. Kaede-Centrin1-labeled centrosome imaged 2 days after photoconversion with mother centriole (red), daughter centriole (green), and stained for a centrosomal protein, γ-tubulin (blue). The overlay image is shown on the right. Scale bar = 2 μm.

Kaede-Centrin1 green

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red

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Before PC

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After PC

Figure 5A.5.5 Fluorescent properties of Kaede-Centrin1 before and after photoconversion (PC). Plasmid DNA encoding Kaede-Centrin1 was injected into the lateral ventricle of an embryo at gestational day 13.5 (E13.5), and the embryo was perfused the following day (E14.5). A 60-μm ßoating brain section was Þrst imaged using a confocal microscope in green (488) and red (546) channels. Kaede-Centrin1 expression is detected in the ventricular zone (VZ) and subventricular zone (SVZ) in the green channel only. Following a 15-sec exposure to unÞltered white light from a mercury lamp, the section was imaged a second time with the same laser output and detection parameters. Note the complete conversion of Kaede-Centrin1 from green- to red-ßuorescent. Scale bar = 10 μm. Current Protocols in Stem Cell Biology

the developing brain, and this segregation has been shown to be critical for maintaining the pool of neural progenitors (Wang et al., 2009). The asymmetric segregation of mother versus daughter centrosomes has been previously reported in other organisms such as Drosophila (reviewed in Yamashita and Fuller, 2008), suggesting that the regulation of centrosome inheritance may be a conserved mechanism for ensuring the proper balance between progenitor maintenance and differentiation during asymmetric cell division in eukaryotes. The Kaede-Centrin1 pulse-labeling and photoconversion protocol distinguishes between mother and daughter centrosomes, allowing a closer study of this mechanism in real-time in the developing mammalian brain (Fig. 5A.5.4).

Critical Parameters and Troubleshooting The combination of in utero electroporation followed by in utero photoconversion is

stressful for the mouse and can result in a high rate of embryo mortality. There are a few key points to heed when performing these procedures that can increase the probability of embryo survival. In general, the surgeries for in utero injection and electroporation and photoconversion should be performed in the minimum amount of time allowed by the experimenter’s skills. The embryos, as well as the mother’s internal organs and overlying abdominal muscle and skin, should be kept moist (with warm PBS) at all times. For Basic Protocol 2, the time of exposure to violet light is the critical parameter. If the time is too short, effective photoconversion cannot take place; however, too long an exposure of the embryos to the outer environment can substantially decrease viability. It is worth noting that with repeated use, the intensity of light from a mercury lamp decreases. If possible, a bulb dedicated to

red

Kaede-Centrin1 green

overlay

1 1

Pia

NCX 2

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Figure 5A.5.6 Segregation of mother and daughter centrosomes in the developing neocortex after pulse-labeling with Kaede-Centrin1. Embryonic neocortex (NCX) at gestational day 16.5 (E16.5) treated with a nuclear stain. The ganglionic eminence (GE) and pial surface (Pia) are indicated as points of reference. Scale bar = 200 μm. Plasmid DNA expressing Kaede-Centrin1 was injected into the lateral ventricle of an embryo at E13.5, and the embryo was exposed in utero to violet light at E14.5 to effect photoconversion of the Kaede-Centrin1 protein from green to red. The embryo was perfused at E16.5, and a 60-μm ßoating brain section was imaged by confocal microscopy. Mother centrosomes retain red ßuorescence from the photoconversion, and are primarily localized in the ventricular zone (VZ), the cortical progenitor niche (2). Daughter centrosomes, born after the pulse of violet light was administered, are green only, and are found in both the VZ as well as in the cortical plate (1), where differentiated neurons have migrated. Note the absence of red ßuorescence signal in this area. Scale bar = 10 μm.

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photoconversion should be set aside to be used only for this purpose, so as to keep track of the burn use time. In addition, to maximize photoconversion, the embryos should be positioned under the light beam such that the beam covers the entire surface of the cortex on the injected side of the brain. Finally, Kaede is a highly photolabile protein that is easily converted from green to red ßuorescent by prolonged exposure to ambient levels of UV light or a brief exposure to (unÞltered) white light from a mercury lamp (Fig. 5A.5.5). This is true even for Þxed tissue. Thus, care must be taken during processing and handling of the tissue and subsequent imaging, in order to protect the tissue from any unwanted photoconversion.

Anticipated Results Centrosome duplication is initiated at the G1 /S transition and completed before mitosis, and an additional 1.5 to 2 cell divisions are required for the nascent centriole to fully mature (Delattre and Gonczy, 2004; Anderson and Stearns, 2009). During the peak phase of neurogenesis in mice (E13.5 to 16.5), the cell-cycle length is ∼12 to 18 hr (Mitsuhashi and Takahashi, 2009). Therefore, the segregation of mother centrosome- and daughter centrosome-containing cells can be observed in the period of ∼36 to 48 hr after photoconversion. If a complete photoconversion of Kaede protein takes place, one expects that green and red ßuorescent centrosomes (i.e., mother centrosomes) remain primarily at the ventricular zone surface where radial glia reside, while centrosomes with green ßuorescence only (i.e., daughter centrosomes) populate the more dorsal layers of the neocortex where the differentiated progeny of radial glia have migrated (Fig. 5A.5.6). Note the following points: a. There is a mosaic incorporation of plasmid DNA in the area of electroporation, therefore, not all radial glia will be labeled. b. If performed correctly, all electroporated embryos should take up plasmid DNA. c. EfÞciency of photoconversion should be 100%, as determined by pilot experiments. d. Labeled centrosomes should be visible by epißuorescence at 40× or 60× objective if experiment is performed as described.

Acknowledgements We thank Drs. She Chen and Shuijin He for technical assistance in the preparation of this manuscript, and Drs. Tsai Jin-Wu and Richard B. Vallee for help with ex vivo photoconversion and time-lapse imaging studies. Research in the Shi laboratory is supported by the Whitehall Foundation, March of Dimes, NARSAD, Klingenstein Foundation, Dana Foundation, Autism Speaks, NIDA, and NIMH.

Literature Cited Anderson, C.T. and Stearns, T. 2009. Centriole age underlies asynchronous primary cilium growth in mammalian cells. Curr. Biol. 19:1498-1502. Daza, R.A.M., Englund, C., and Hevner, R.F. 2007. Organotypic slice culture of embryonic brain tissue. Cold Spring Harbor Protoc. doi:10.1101/pdb.prot4914. Delattre, M. and Gonczy, P. 2004. The arithmetic of centrosome biogenesis. J. Cell Sci. 117:16191630. Elias, L. and Kriegstein, A. 2007. Organotypic slice culture of E18 rat brains. J. Vis. Exp. 6:235. G¨otz, M. and Huttner, W.B. 2005. The cell biology of neurogenesis. Nat. Rev. Mol. Cell Biol. 6:777788. Mitsuhashi, T. and Takahashi, T. 2009. Genetic regulation of proliferation/differentiation characteristics of neural progenitor cells in the developing neocortex. Brain Dev. 31:553-557. Molyneaux, B.J., Arlotta, P., Menezes, J.R., and Macklis, J.D. 2007. Neuronal subtype speciÞcation in the cerebral cortex. Nat. Rev. Neurosci. 8:427-437. Nigg, E.A. and Raff, J.W. 2009. Centrioles, centrosomes, and cilia in health and disease. Cell 139:663-678. Schatten, H. and Sun, Q.Y. 2010. The role of centrosomes in fertilization, cell division and establishment of asymmetry during embryo development. Semin. Cell Dev. Biol. 21:174184. Tabata, H. and Nakajima, K. 2008. Labeling embryonic mouse central nervous system cells by in utero electroporation. Dev. Growth Differ. 50:507-511. Wang, X., Tsai, J.W., Imai, J.H., Lian, W.N., Vallee, R.B., and Shi, S.-H. 2009. Asymmetric centrosome inheritance maintains neural progenitors in the neocortex. Nature 461:947-955. Yamashita, Y.M. and Fuller, M.T. 2008. Asymmetric centrosome behavior and the mechanisms of stem cell division. J. Cell Biol. 180:261-266.

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Generation of Human Embryonic Stem Cell Reporter Knock-In Lines by Homologous Recombination

UNIT 5B.1

Richard P. Davis,1,2 Catarina Grandela,1,3 Koula Sourris,1 Tanya Hatzistavrou,1 Mirella Dottori,4 Andrew G. Elefanty,1 Edouard G. Stanley,1 and Magdaline Costa1 1

Monash Immunology and Stem Cell Laboratories, Monash University, Clayton, Australia Department of Anatomy and Embryology, Leiden University Medical Centre, Leiden, The Netherlands 3 Laboratory of Experimental Oncology and Radiobiology (LEXOR), Center for Experimental Molecular Medicine, Academic Medical Center, The Netherlands 4 Centre for Neuroscience and Dept of Pharmacology, The University of Melbourne, Parkville, Australia 2

ABSTRACT This unit describes a series of technical procedures to form clonal human embryonic stem cell (hESC) lines that are genetically modified by homologous recombination. To develop a reporter knock-in hESC line, a vector is configured to contain a reporter gene adjacent to a positive selection cassette. These core elements are flanked by homologous sequences that, following electroporation into hESCs, promote the integration of the vector into the appropriate genomic locus. The positive selection cassette facilitates the enrichment and isolation of genetically modified hESC colonies that are then screened by PCR to identify correctly targeted lines. The selection cassette, flanked by loxP sites, is subsequently excised from the positively targeted hESCs via the transient expression of Cre recombinase. This is necessary because the continued presence of the cassette may interfere with the regulation of the reporter or neighboring genes. Finally, these genetically modified hESCs are clonally isolated using single-cell deposition flow cytometry. Reporter knock-in hESC lines are valuable tools that allow easy and rapid identification and isolation of specific hESC derivatives. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 11:5B.1.1-5B.1.34.  Keywords: human embryonic stem cells r hESCs r gene targeting r homologous recombination r fluorescent reporter gene

INTRODUCTION This unit describes a series of procedures to form clonal genetically modified human embryonic stem cell (hESC) lines in which DNA sequences encoding fluorescent or other reporter genes are inserted into the genome by homologous recombination. Consequently, the reporter gene mirrors the expression pattern of the endogenous gene it replaces because it is regulated by the same transcriptional mechanisms. Targeting of reporter genes into the loci of lineage-specific transcription factors has facilitated the isolation and enrichment of specific cell types from differentiating mouse and human embryonic stem cells that would otherwise be unobtainable (Ying et al., 2003; Fehling et al., 2003; Ng et al., 2005; Micallef et al., 2005, 2007; Gadue et al., 2006; Davis et al., 2008a). A combination of molecular biology and specialized cell culturing techniques are required to create a reporter knock-in hESC line (Fig. 5B.1.1). The gene-targeting vector is a key element in these procedures, and factors to be taken into consideration when constructing

Current Protocols in Stem Cell Biology 5B.1.1-5B.1.34 Published online November 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05b01s11 C 2009 John Wiley & Sons, Inc. Copyright 

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Enzymatically expand hESCs (UNIT 1C.1)

Design and construct a targeting vector (Strategic Planning)

Electroporate hESCs with targeting vector and select for stably transfected cells (Basic Protocol 1)

Following selection, isolate and expand the hESC colonies (Support Protocol 1)

Extract DNA and screen by PCR for targeted hESCs (Support Protocols 2 and 3)

No

Is there a correctly targeted hESC colony? Yes

Transfer the targeted hESC colony into organ culture dishes and expand enzymatically (UNIT 1C.1)

Confirm: 1) targeting by Southern blot analysis or DNA sequencing; 2) stem cell phenotype (UNITS 1B.3 and 1B.4); 3) normal karyotype.

Remove the positive selection cassette from hESCs by transient transfection with the Cre-expression vector. Select for transfected hESCs (Basic Protocol 2)

Isolate and expand the hESCs colonies (Support Protocol 1)

Extract DNA and screen by PCR for loss of the selection cassette and non-integration of Cre-expression plasmid (Support Protocols 2 and 3)

No

Does a hESC colony satisfy the above requirements? Yes

Figure 5B.1.1

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Transfer hESC colony into organ culture dishes and expand into bulk culture (UNIT 1C.1)

Clonally derive sublines by single-cell deposition flow cytometry (Support Protocol 4)

Isolate and expand the resulting hESCs colonies (Support Protocol 1)

Extract DNA and screen by PCR to confirm line is targeted, has lost the positive selection cassette, and there is non-integration of Cre-expression plasmid (Support Protocols 2 and 3)

Does a hESC colony satisfy the above requirements?

No

Yes

Maintain and expand the hESC line (UNIT 1C.1)

Transfer the reporter hESC line into organ culture dishes (UNIT1C.1)

Confirm: 1) targeting by Southern blot analysis or DNA sequencing; 2) stem cell phenotype (UNITS 1B.3 and 1B.4); 3) normal karyotype.

Cryopreserve stocks of the hESC line

Determine the fidelity of the reporter gene expression

Figure 5B.1.1 (continued) A schematic representation of the sequence of procedures to generate a clonal reporter knock-in hESC line in which the positive selection cassette has been removed from the genome. Where possible, reference is made to either the relevant protocol or section in this unit to perform the step, or to another appropriate unit.

the targeting vector are detailed in the Strategic Planning section. This section also describes methods for maintaining and enzymatically expanding hESCs. The stable integration of the targeting vector into hESCs by electroporation and the subsequent isolation of these cells by drug selection are described in Basic Protocol 1. Support Protocol 1 outlines the process involved in picking and replicating the drugresistant hESC colonies, and this is followed by methods detailing the extraction of DNA from the colonies (Support Protocol 2 or the Alternate Protocol). A PCR screening strategy is then employed to identify the gene-targeted hESC colonies (Support Protocol 3). The new hESC lines that contain the correct genetic modification are maintained as mechanically passaged colonies (UNIT 1C.1), and are also expanded enzymatically for the deletion of the positive selection cassette as described (UNIT 1C.1).

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Elements located within the positive selection cassette can interfere with the expression of both the reporter gene and neighboring endogenous genes (Hug et al., 1996; Pham et al., 1996; Scacheri et al., 2001; R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.). Therefore, a procedure for a Cre recombinase–mediated excision of the positive selection cassette from the targeted hESC lines is detailed in Basic Protocol 2. Finally, Support Protocol 4 describes a method for the clonal isolation and expansion of these gene-targeted hESCs using the single-cell deposition function of a flow cytometer. These protocols were employed to generate multiple independent MIXL1GFP/w hESC lines (Davis et al., 2008a). The expression of green fluorescent protein (GFP) during the directed differentiation of these hESCs enabled cells expressing the transcription factor MIXL1 to be identified and isolated. These techniques have also been successfully used to generate reporter knock-in hESC lines targeted at nine additional loci (Costa et al., 2007; A.G. Elefanty and E.G. Stanley, unpub. observ.).

STRATEGIC PLANNING Design of the Targeting Vector This section outlines some of the general principles to consider when designing the gene-targeting vector. A detailed description of the techniques and methods used to engineer a targeting vector are beyond the scope of this unit, but are available in other publications (Struhl, 2001; Elion et al., 2007; Thomason et al., 2007). The standard constituents of a targeting vector for the generation of a reporter knock-in hESC line include a promoterless reporter gene followed by a positive selection cassette comprising of a constitutively active promoter that regulates the expression of an antibiotic-resistance gene. These core elements are flanked by two arms that are homologous to the target locus (Fig. 5B.1.2). Given that the gene-targeting vectors are routinely introduced into ESCs by electroporation in a linearized form, at least one unique restriction enzyme site is located outside the homologous sequences. This is typically at the junction of the homologous arms and the plasmid backbone. The reporter gene is typically a fluorescent marker, such as GFP or red fluorescent protein (RFP), which lacks a 5 -untranslated region and an ATG start codon. Therefore, the coding sequence of the reporter gene is placed in frame with the initiation codon of the target gene. This measure improves the likelihood of recapitulating the expression profile of the endogenous gene. Targeting the reporter gene to the 5 end of the coding sequence also ensures that no wild-type polypeptide is translated upstream to the reporter gene, resulting in the deliberate ablation of expression of that allele and preventing the creation of a dominant negative allele. The sequences inserted into the genome generally replace a minimal portion of an exon to avoid the disruption of the RNA splicing sequences and the loss of introns. It is also recommended to avoid targeting into alternatively spliced exons.

Generation of hESC Reporter Knock-In Lines

While the optimal length of the homologous sequences for gene targeting in hESCs has not been thoroughly investigated, the inclusion of one homology arm greater than 6 kb improves the targeting frequency (Zwaka and Thomson, 2003; Costa et al., 2007). Targeting vectors generally contain one long and one short homology arm. The shorter homology arm must be long enough to facilitate recombination, but short enough to screen stable transfectants for recombination events by PCR. In practice, we have found that this homology arm should lie between 2 and 4 kb in size, while the combined lengths of the two arms should be 10 to 14 kb in size. Although longer homology arms can improve the targeting frequency, vectors >18 kb in length can limit both the propagation of standard bacterial strains and the repertoire of unique restriction enzyme sites available to linearize the vector prior to electroporation.

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pos se

lect .

3’

ar m

orter rep

ya

5’ h o

mo log y

log mo ho

*

*

neg . sele ct.

rm

vector backbone

Figure 5B.1.2 Plasmid map describing the possible structure of the targeting vector to generate a knock-in hESC line by homologous recombination. Two genomic fragments (5 homology arm and 3 homology arm) flank a reporter gene (green arrow) lacking an initiation codon, and a positive selection cassette (blue rectangle). The reporter gene is placed in frame with the ATG start codon included in the 5 homology arm. The red triangles represent loxP sequences that flank the positive selection cassette. A negative selection cassette (purple rectangle) can be included in the construct if desired. Unique restriction sites (marked as *) are located within the plasmid backbone to linearize the targeting construct prior to electroporation.

The homology arm sequences may also affect the success of homologous recombination. While targeting vectors containing nonisogenic sequences decrease the frequency of homologous recombination up to 20-fold in mESCs (te Riele et al., 1992; van Deursen and Wieringa, 1992), the efficiency of gene targeting in hESCs appears similar regardless of the origin of the homology arms. Practically, this also means that the same knock-in vector will target a given locus in different hESC lines at similar frequencies (Costa et al., 2007). Following recombination, positive selection cassettes are required to permit identification of stably transfected hESCs, which occur at a frequency between 1 in 104 105 electroporated cells. The neomycin phosphotransferase gene (neo) is highly expressed in hESCs when regulated by the mouse phosphoglycerate kinase (PGK) promoter (R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.), and this selection cassette is routinely used by the authors when transfecting hESCs. Additionally, if the hESCs are being cultured on mouse embryonic fibroblast feeder cells, geneticin (G418)-resistant mouse lines are readily available. As an alternative, the neo gene may also be replaced with the hygromycin B phosphotransferase (hph) gene and the antibiotic hygromycin B used to enrich for stably transfected hESCs (L. Azzola, A.G. Elefanty, and E.G. Stanley, unpub. observ.). In correctly targeted clones, retention of the positive selection cassette in the genome can influence expression of the target locus and of neighboring genes (Hug et al., 1996; Pham et al., 1996; Scacheri et al., 2001; R. Davis, A.G. Elefanty, and E.G. Stanley, unpublished observations). Therefore

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flanking the cassette with loxP sequences allows for subsequent removal by transient expression of Cre recombinase (Gu et al., 1993). Replacement-type targeting vectors can also include a negative selection cassette, such as the herpes simplex virus thymidine kinase (HSVTK) gene, to allow for selection against random integrants and enrich for targeted recombinants (Zwaka and Thomson, 2003). This cassette is located outside the region of homology to the target gene, normally at the end of the short homology arm.

Culturing the hESCs The adaptation of the hESCs to enzymatic passaging using either trypsin or TrypLE Select (Invitrogen) is essential for success in the protocols described. Without such preconditioning, disaggregation of hESCs to a single-cell suspension results in extensive cell death (UNIT 1C.1). Protocols for the adaptation and expansion of enzymatically passaged hESCs are described in detail in UNIT 1C.1 and UNIT 1D.3. Only hESCs that have undergone 5 to 10 enzymatic passages should be used in Basic Protocols 1 and 2, and Support Protocol 4. Validation of hESC Reporter Knock-in Lines The protocols in this unit describe the generation of hESC reporter lines by homologous recombination, identification of correctly targeted clones, removal of the selectable marker, and single-cell cloning of the cell line. However, before any hESC reporter line is used for experimentation, further screening procedures are required to validate the cell line. Southern blot analysis of the putatively targeted hESC clones can confirm that the desired recombination events have occurred at both the 5 and 3 ends, and that the cells contain only a single copy of the reporter gene. A detailed description of Southern blotting is outside the scope of this unit. In brief, genomic DNA extracted from the hESC colonies is digested with restriction enzymes, electrophoresed on an agarose gel, and transferred to a membrane. This membrane is hybridized with radiolabeled probes to the gene locus that lies external to the region of homology with the targeting vector, and the labeled DNA fragments detected using either a phosphoimager or X-ray film. Such probes serve to identify correctly targeted alleles by virtue of size differences between hybridizing fragments generated by wild-type and targeted alleles. These size differences arise because of the incorporation of additional DNA sequences and/or new restriction enzyme sites associated with the introduction of the reporter gene and selectable marker sequences into the native locus. In addition, a radiolabeled probe that hybridizes with the coding sequence of the reporter gene can confirm that only a single integration event occurred. The genetically manipulated hESC line must also maintain the characteristics of a stem cell. The cells should be regularly analyzed by flow cytometry for the expression of a panel of stem cell markers including transcription factors, such as OCT4 and NANOG, and the cell surface antigens, such as SSEA3 and 4, Tra-1-60, Tra-1-81, and CD9 (UNIT 1B.3). Additionally, when the genetically modified hESCs are injected into the testes of immunodeficient mice, they should form multi-lineage teratomas (UNIT 1B.4). The karyotype should also be periodically checked to confirm that no karyotypic abnormalities have been introduced during the genetic manipulations. It is recommended that this analysis, to at least the level of G-banding, be performed by a clinical cytogenetics facility. Generation of hESC Reporter Knock-In Lines

NOTE: The following protocols are performed in either a Class I (laminar-flow) biosafety cabinet or a Class II biohazard hood.

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NOTE: All materials and reagents that come into contact with live cells must be sterile and proper aseptic technique must be used when handling the cells or setting up experiments. NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator, unless otherwise specified.

ELECTROPORATION OF hESCs AND SELECTION OF ANTIBIOTIC-RESISTANT hESCs

BASIC PROTOCOL 1

Electroporation is the most common means of generating targeted mESC lines (Giudice and Trounson, 2008). While the number of reports describing gene targeting by homologous recombination in hESCs is currently limited, the majority of these have also utilized electroporation (Zwaka and Thomson, 2003; Costa et al., 2007; Irion et al., 2007; Davis et al., 2008a; Braam et al., 2008). This protocol describes the procedure for electroporating hESCs with a linearized gene-targeting vector, followed by the positive selection of stably transfected colonies. It is recommended that the neo gene be included in the positive selection cassette, allowing for the drug geneticin (G418) to be used as the selection agent.

Materials hESCs in 150-cm2 tissue culture flasks at enzymatic passage 5 to 10 (see UNIT 1C.1) in hESC medium (see recipe) 150-cm2 gelatinized tissue culture flasks (see recipe) preseeded with mitotically inactivated MEFs at 1 × 104 /cm2 for passaging hESCs prior to electroporation MEF medium (see recipe) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) hESC medium (see recipe), 37◦ C Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) 0.4% (w/v) Trypan blue (Fluka) Soybean trypsin inhibitor (see recipe; Invitrogen), optional Linearized targeting vector (see Strategic Planning) in Tris/EDTA (TE) buffer (see recipe) for transfection 60-mm gelatinized tissue culture dishes preseeded with 2 × 104 /cm2 mitotically inactivated MEFs Geneticin/G418 Selective Antibiotic (Invitrogen) Mitotically inactivated, irradiation-treated (UNIT 1C.3) G418-resistant mouse embryonic fibroblasts (MEFs; Conner, 2000) 37◦ C water bath Gene Pulser cuvette, 0.4-cm electrode gap, sterile (Bio-Rad, cat. no. 165-2088) 15- and 50-ml sterile centrifuge tubes Refrigerated centrifuge Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Tissue culture microscope with phase contrast objectives and phase rings Hemacytometer (Neubauer) Electroporator (Gene Pulser II System; Bio-Rad) Sterile Pasteur pipets Prepare hESCs prior to electroporation (day −1, day 0) 1. On the day prior to electroporation, enzymatically dissociate the hESCs from two 150-cm2 flasks and replate onto two fresh 150-cm2 flasks preseeded with MEFs at low density (1 × 104 MEFs/cm2 in 20 ml MEF medium), using either 2 ml trypsin or 2 ml TrypLE Select per 150-cm2 flask (Fig. 5B.1.3A). It is critical that hESCs are adapted to enzymatic passaging using either trypsin or TrypLE Select prior to beginning this protocol. The enzymatic expansion of hESCs from

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A

B

C

D

Figure 5B.1.3 Photomicrographs of hESCs expanded in bulk culture, after electroporation, during drug selection, and following selection. Some groups of hESCs or individual colonies in panels (A), (B), and (C) are outlined by black-dotted lines. (A) hESCs in bulk culture grown on MEFs at reduced density on day of electroporation. (B) hESCs on the day following electroporation. (C) hESCs 5 days after electroporation, just prior to antibiotic selection. (D) A hESC colony following 5 days of selection. (A) through (D) Original magnification 50×.

stock cultures maintained by mechanical passaging is described in detail in UNIT 1C.1. The cells should be enzymatically disaggregated every 3 or 4 days, and split no more than at a ratio of 1:2. Generally, after the first 4 or 5 enzymatic passages, the hESCs will have been expanded into two 150-cm2 flasks. The hESCs are passaged onto fresh MEFs at a ratio of 1:1 on the day prior to the electroporation to ensure that the cells are actively dividing and to remove dead and dying cells. Seeding the flasks with a reduced density of MEFs (1 × 104 MEFs/cm2 compared with 2 × 104 MEFs/cm2 ), results in a partial depletion of feeder cells.

2. At a time point ∼2 to 3 hr before the electroporation, aspirate the medium and re-feed the cells with fresh hESC medium. 3. At a time point ∼2 hr before the electroporation, place a 10-ml aliquot of CMF-PBS on ice and a 40-ml aliquot of hESC medium in a water bath at 37◦ C. 4. Place the Gene Pulser electroporation cuvette on ice.

Harvest the cells 5. Harvest the hESCs passaged the day before. First, aspirate the hESC medium and then rinse the flasks with 5 to 10 ml CMF-PBS. 6. Add 2 ml of trypsin or TrypLE Select to each 150-cm2 flask and ensure that the dissociation solution coats the surface of the cells. Incubate 4 min at 37◦ C in the incubator. Check that the hESCs dislodge from the flasks with gentle tapping. Generation of hESC Reporter Knock-In Lines

If the cells are dissociated with trypsin, then a neutralization step is required. Add 1 ml of soybean trypsin inhibitor to each flask and swirl to mix. A specific neutralizing agent is not required for TrypLE Select.

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7. Add 8 ml of hESC medium to each flask, mix, and transfer the contents of the two flasks to a single 50-ml centrifuge tube. 8. Pellet the cells by centrifuging the tube 3 min at 480 × g, at 4◦ C, and aspirate the supernatant. 9. Resuspend the hESCs in 10 ml of CMF-PBS and transfer the cell suspension to a 15-ml centrifuge tube. 10. Using a Gilson pipettor, mix 10 μl of the cell suspension with 10 μl trypan blue. Load 10 μl onto a hemacytometer and, using a tissue culture microscope, perform a cell count (UNIT 1C.3). Subtract the total MEF number (3 × 106 ) from the count. A total of 1 × 107 hESCs is required per electroporation. Two 150-cm2 flasks seeded with MEFs at a density of 1 × 104 /cm2 will contain ∼3 × 106 MEFs. After subtracting the MEF count, a semi-confluent 150-cm2 flask of hESCs typically contains between 6–8 × 106 hESCs. Therefore the yield from two such flasks (∼12–16 × 106 hESCs) will provide enough hESCs for a single electroporation (∼10 × 106 hESCs). The remaining hESCs can be replated onto a new flask seeded with MEFs at the regular density (2 × 106 MEFs/cm2 ) to maintain the undifferentiated culture. If there are ∼2–4 × 106 hESCs remaining, these should be plated onto a 75-cm2 flask, while a 150-cm2 flask should be used if there are in excess of 4 × 106 hESCs.

Prepare the cells with the targeting vector for electroporation 11. Centrifuge the cells again for 3 min at 480 × g, 4◦ C, aspirate the supernatant, and resuspend 1 × 107 hESCs in a final volume of 750 μl ice-cold CMF-PBS. 12. Using a Gilson pipettor, add 50 μl of TE buffer containing between 10 and 20 μg of the linearized gene-targeting vector to the electroporation cuvette. 13. Carefully transfer the hESC suspension into the cuvette using a pipettor, ensuring that the DNA-containing TE buffer and hESCs are evenly resuspended. 14. Place the cuvette containing the DNA and cell suspension mix on ice for 5 min.

Electroporate the cells 15. Wipe the outside of the cuvette to remove any water or ice before electroporating the cells at 250 V and 500 μF (Costa et al., 2007). Other groups have achieved successful transfection of the plasmid by electroporating the hESCs at 320 V and 200 μF (Zwaka and Thomson, 2003), or 320 V and 250 μF (Braam et al., 2008). In our laboratory, our conditions have been used to target at least 9 loci (Costa et al., 2007; A.G. Elefanty and E.G. Stanley, unpub. observ.).

Plate the electroporated cells 16. Using a Pasteur pipet, transfer the contents of the cuvette to a 50-ml centrifuge tube containing 10 ml of prewarmed hESC medium. 17. Centrifuge the electroporated cells 3 min at 480 × g, room temperature (20◦ to 25◦ C). 18. Carefully aspirate the supernatant and gently resuspend the pellet in 15 to 18 ml of prewarmed hESC medium. Steps 16 to 18 remove cellular debris that could impair the viability of the surviving hESCs.

19. Plate the cell suspension into five or six 60-mm dishes preseeded with 2 × 104 MEFs/cm2 , and incubate the dishes in a humidified incubator at 37◦ C, 5% CO2 (Fig. 5B.1.3B).

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The number of 60-mm dishes that the electroporated hESCs are plated into is dependent on the proportion of hESCs that survive the electroporation procedure. This percentage not only varies between hESC lines, but also reflects the degree to which the cells have adapted to the enzymatic passaging prior to electroporation. Plating the hESCs into five or six 60-mm dishes usually ensures the hESC density is low enough that the cells can be left to recover and proliferate for 4 to 5 days before beginning selection. If the hESC cultures are confluent in less than 4 days, plate the cells into a larger number of 60-mm dishes when performing future electroporations.

20. Two days after the electroporation, gently aspirate the medium containing the dead cells from each dish and replace with 4 ml of fresh hESC medium per dish. Repeat this daily.

Select for geneticin-resistant hESCs (day 4 or 5) 21. Supplement 200 ml of hESC medium with the drug geneticin (G418) so that the final concentration is 50 μg/ml. The recommended final concentration of G418 to use is 50 μg/ml; however, the optimal concentration for selection may vary for different hESC lines. Prior to performing an electroporation, determine the minimum concentration of G418 required to eliminate G418-sensitive hESCs within 5 days of addition. For 1 week of selection, 200 ml of hESC medium containing G418 should be sufficient stock. This medium has a limited lifespan of 7 days.

22. Apply 50 μg/ml G418 selection to the hESCs when the dishes are ∼60% to 80% confluent with hESCs (4 or 5 days following electroporation; Fig. 5B.1.3C). A 60-mm dish that is semi-confluent with nontransfected hESCs should also be placed under selection as a control.

23. Change the selection medium daily. Frequent medium changes are necessary during selection to remove the dead cells.

24. Four days into selection, supplement the dishes with fresh MEFs at 1 × 104 MEFs/cm2 . Resuspend the required number of MEFs in 20 ml hESC medium containing G418 and dispense evenly over all the dishes, such that each dish contains a total volume of 4 ml. MEFs can support hESC growth for ∼1 week, and so the dishes will need to be supplemented with additional MEFs during the protocol to maintain densities of ∼2 × 104 MEFs/cm2 . This should be done either 8 days after the initial plating of the electroporated hESCs (Fig. 5B.1.3D), or if the MEF density appears low after the onset of selection. Supplementation with fresh MEFs is required even if the MEFs are G418 resistant.

Remove the selection agent 25. Following 7 days of G418 selection, return to culturing the G418-resistant cells in hESC medium without G418. After 7 days, the control 60-mm dish should contain no residual viable hESCs. If live hESCs remain on the control dish, continue G418 selection on all dishes until colony transfer (∼18 days post electroporation). Alternatively, the dose of G418 required to kill control hESCs may need to be re-titrated.

26. Allow the colonies to grow for approximately a further 7 days, changing the hESC medium daily. The colonies are ready to transfer to 48-well tissue culture plates (Support Protocol 1) when they are ∼2 mm in diameter. If the cells begin to differentiate, they should be transferred sooner. Generation of hESC Reporter Knock-In Lines

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PICKING AND EXPANDING ANTIBIOTIC-RESISTANT hESC COLONIES This protocol describes transferring and expanding individual genetically modified hESC colonies that emerge on the 60-mm dishes following the selection procedures described in Basic Protocols 1 and 2. Approximately 7 to 9 days after the withdrawal of the selection agent from the culture medium, each hESC colony is mechanically passaged into a well of a 48-well tissue culture plate (Fig. 5B.1.4A,B). In our experience, the number of antibiotic-resistant colonies after Basic Protocol 1 may vary from 50 to more than 300 per 107 electroporated hESCs. One week after the initial transfer, the hESCcontaining plates are then duplicated, with one plate destined for extraction of DNA to perform the PCR screen (Support Protocols 2 and 3), while the other plate will serve to maintain the hESC colonies in culture until positive clones are identified.

SUPPORT PROTOCOL 1

Materials Flat-bottomed 48-well tissue culture plates, gelatinized (see recipe) and preseeded with mitotically inactivated MEFs at 2 × 104 /cm2 Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) hESC medium (see recipe) 60-mm tissue culture dishes containing drug-resistant hESC colonies from either Basic Protocol 1 or 2 26-G, 1/2 -in. (0.45 × 13–mm) needles 1-ml syringe 200-μl pipet tips Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Stereomicroscope Prepare 48-well tissue culture plates for transfer of the hESC colonies (day −1) 1. Estimate the total number of individual, undifferentiated hESC colonies to be picked from the tissue culture dishes. Seed enough 48-well plates with ∼0.75 × 106 (MEFs in 12 ml MEF medium, 250 μl/well) MEFs/plate, such that each hESC colony can be transferred to an individual well. These 48-well plates are designated the “Primary” plates.

Figure 5B.1.4 Photomicrographs of a hESC colony being transferred and expanded. (A) 25× magnification of a hESC colony sliced into a grid pattern prior to dislodgement and transfer to a well on a 48-well plate. (B) Transferred pieces from a single hESC colony grown for 7 days in a well on a 48-well plate.

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It is recommended that one person should not pick more than 150 hESC colonies at a time due to the difficulty in maintaining and processing large numbers of colonies later in the PCR screening (Support Protocol 3). If the electroporation performed in Basic Protocol 1 has been particularly successful, a pair of researchers can manage to pick, replicate, and process a larger number of antibiotic-resistant colonies.

Pick hESC colonies (day 0) 2. Aspirate the MEF medium from the 48-well plates pre-seeded with MEFs, and dispense 200 μl fresh hESC medium into each well. 3. Aspirate the hESC medium from the 60-mm dishes containing the hESC colonies and supplement with 4 ml of fresh hESC medium. 4. Using a 26-G needle attached to a 1-ml syringe, cut a hESC colony in a grid motif to generate a minimum of 16 small pieces (Fig. 5B.1.4A). Detach the pieces from the dish by flicking them off with the needle, or with a 200-μl pipet tip attached to a pipettor. These procedures are performed under a zoom-focus stereomicroscope.

5. Using a 200-μl pipet, collect all the pieces and transfer them into a well on one of the “Primary” 48-well plates. 6. Discard the 26-G needle and 200-μl pipet tip after harvesting the hESC colony. 7. Repeat steps 4 to 6 to harvest the remaining hESC colonies on the tissue culture dishes. Select well-spaced colonies to avoid cross-contamination of clones. Avoid picking colonies that are significantly smaller than the majority of the colonies. Smaller colonies can be left to expand longer on the 60-mm plates and picked several days later when larger.

8. Replace the hESC medium in the wells daily with 200 μl of fresh medium per well. Within 2 days of picking, the hESCs should be visible with the formation of multiple colonies in each well.

Replicate the hESC-containing 48-well plates (days 5 to 7) 9. Between 5 and 7 days after picking the hESC colonies, verify that the wells are approaching confluence (Fig. 5B.1.4B). Prepare two sets of 48-well plates with MEFs (as described in step 1). Label one set of plates “DNA,” and the other set “Maintenance.” 10. The next day, replace the MEF medium on the multi-well plates labeled “DNA” and “Maintenance” with 100 μl of fresh hESC medium. 11. Replace the hESC medium on the 48-well plates labeled “Primary” with 300 μl of hESC medium. 12. Using a plugged 200-μl pipet tip attached to a pipettor, scrape the bottom of a well on a “Primary” plate in a criss-cross fashion until the hESC colonies have detached. Examine the well under a stereomicroscope to confirm that the hESC colonies have broken into small cell clumps. If the hESCs have not fragmented into small clumps, pipet the cells up and down to try to break up the colonies.

Generation of hESC Reporter Knock-In Lines

13. Transfer 100 μl of the well contents to a 48-well plate labeled “Maintenance,” and the remaining 200 μl to the corresponding well on the plate labeled “DNA.” Expel air bubbles into the medium to assist in identifying wells that contain passaged hESCs. Discard the 200-μl pipet tip after use.

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Uneven distribution of the hESC clones between the 48-well plates allows the plates labeled “DNA” to be processed and screened by PCR before the hESCs on the “Maintenance” plates have reached confluence and need to be passaged. If the hESCs in the wells of the “Primary” plate are confluent, and if there are fewer than 48 hESC clones to screen, it is often convenient to directly isolate DNA for PCR screening at this stage without further expanding the cell numbers. Instead of seeding the cells into the wells of the “DNA” plate, transfer the hESC clones into individually labeled microcentrifuge tubes, and pellet the cells by centrifuging 3 min at 480 × g, 4◦ C. The genomic DNA can be isolated using the method described in the Alternate Protocol.

14. Repeat steps 12 and 13 to transfer all remaining hESC colonies. 15. The next day, replace the medium on all plates with 200 μl of fresh hESC medium. Change the hESC medium daily. 16. Approximately 6 days after duplicating the hESC clones, the hESCs on the “DNA”labeled plates will be confluent enough to isolate sufficient DNA for PCR screening (Fig. 5B.1.4B). Proceed to Support Protocol 2 for instructions on how to process these plates. 17. While screening the hESCs to identify clones that contain the correct genetic modification, continue to change the hESC medium daily on the “Maintenance” plates. If the majority of hESC-containing wells on the “Maintenance” plates are more than 80% confluent, or the hESC colonies have begun to differentiate before the PCR screening is complete, the cells will need to be passaged. This is performed as described in steps 9 to 14, but not in duplicate.

PREPARATION OF GENOMIC DNA FROM hESCs GROWING IN 48-WELL TISSUE CULTURE PLATES

SUPPORT PROTOCOL 2

To maximize the quantity of genomic DNA obtained from the genetically modified hESC clones, the cells can be let to overgrow and the plates should only be harvested when the majority of the clones are confluent. This protocol describes a simple, but effective, method of DNA extraction from hESCs growing in 48-well plates. This DNA is then used in a PCR screen to identify hESC colonies that contain the correct genetic modification. In some cases, the PCR screen is not sensitive enough to identify positive clones from DNA isolated using this method. Under these circumstances, the DNA should be extracted from the hESCs incorporating a phenol/chloroform extraction step as described in the Alternate Protocol.

Materials 48-well tissue culture plates labeled “DNA,” containing confluent hESC colonies (from Support Protocol 1, step 13) Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) DNA lysis buffer containing 200 μg/ml proteinase K (see recipe) 100% (v/v) and 70% (v/v) ethanol Tris/EDTA buffer (TE buffer; see recipe) Temperature-adjustable incubator Centrifuge with multi-well plate spinner attachment Blotting paper Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips 1. Aspirate the medium from the plates assigned for DNA extraction and rinse the wells with 200 μl CMF-PBS.

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2. Add 100 μl DNA lysis buffer containing 200 μg/ml proteinase K to each well and incubate 3 hr at 55◦ C. Approximately 5 ml of lysis buffer is required for each plate. Alternatively, the plates can be incubated overnight at 37◦ C.

3. Add 250 μl of 100% ethanol to each well and mix by gently tapping the plate. Mix the solutions until a DNA precipitate is visible in each well.

4. Centrifuge the plates 15 min at 3000 × g for 15 min, room temperature, to pellet the DNA in the bottom of each well. 5. Decant the supernatant by carefully inverting the plate on blotting paper. Take care to ensure that the DNA precipitates remain in the wells.

6. Add 250 μl of 70% ethanol to each well. Centrifuge the plates again 15 min at 3000 × g, room temperature, to ensure the DNA is pelleted on the bottom of the wells. This step removes residual salt from the DNA pellet.

7. Remove the 70% ethanol solution from each well. First, invert the plate on blotting paper and then carefully remove any residual ethanol in each well using a 200-μl pipet with a plugged tip attached. Leave the DNA to air dry for no more than 10 min at room temperature. Use a different 200-μl plugged tip for each well.

8. Resuspend the DNA in 100 μl of TE buffer and incubate for 3 to 4 hr at 55◦ C to dissolve the pellet. The DNA can be stored for at least 3 months at 4◦ C. ALTERNATE PROTOCOL

ISOLATION OF GENOMIC DNA FROM hESCs USING PHENOL/CHLOROFORM EXTRACTION This protocol describes an alternative method of isolating genomic DNA from hESCs grown in multi-well plates. A phenol/chloroform extraction is included to improve the quality of the genomic DNA. This approach should be considered if no hESC clones containing the desired genetic modification are identified when DNA is isolated using the techniques described in Support Protocol 2, or it is already known that the genomic fragment is difficult to amplify in the PCR screening strategy. This protocol can also be used to give high-quality DNA directly from cells on the primary plate if all the wells containing hESCs are confluent, and if there are fewer than 48 clones to screen, saving ∼1 week of culturing time (as described in Support Protocol 1). If there are more than 48 clones, it is difficult for one person to process all of the samples using this procedure.

Materials 48-well tissue culture plates containing confluent hESC colonies (Support Protocol 1, step 13) or microcentrifuge tubes containing clumps of hESCs (Support Protocol 1, step 13, annotation) DNA lysis buffer containing 200 μg/ml proteinase K (see recipe) Phenol/chloroform/isoamyl alcohol (25:24:1) solution saturated with 10 mM Tris·Cl, pH 8.0/1 mM EDTA 70 %(v/v) and 100% (v/v) ethanol Trypsin/EDTA buffer (TE buffer; see recipe) Generation of hESC Reporter Knock-In Lines

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1.5-ml microcentrifuge tubes Vortex mixer Microcentrifuge 55◦ C incubator Current Protocols in Stem Cell Biology

1. Aspirate the medium from the hESC clones growing on the 48-well plates labeled “DNA.” If the hESCs are in microcentrifuge tubes (from Support Protocol 1), pellet the cells by centrifuging 3 min at 480 × g, 4◦ C, before removing the medium.

2. Add 100 μl DNA lysis buffer containing 200 μg/ml proteinase K to each clone and incubate for ∼3 hr at 55◦ C. Alternatively, incubate overnight at 37◦ C.

3. Transfer the contents of each well to individual 1.5-ml microcentrifuge tubes. Label the tubes such that the corresponding hESC clone can be identified on the “Maintenance” multi-well plate. If the hESCs are already in microcentrifuge tubes, skip step 3 and proceed to step 4.

4. Add an equal volume of phenol/chloroform/isoamyl alcohol solution to each tube, vortex to mix well, and then centrifuge the tubes 10 min at 10,000 × g, room temperature. 5. Carefully transfer the top layer containing the DNA to a new 1.5-ml microcentrifuge tube containing 250 μl of 100% ethanol. Mix the solutions until the DNA precipitates. 6. Pellet the DNA by microcentrifuging 10 min at maximum speed, room temperature. 7. Carefully remove the supernatant and wash the DNA pellet with 250 μl of 70% ethanol. Ensure all ethanol is removed by leaving the DNA pellet to air dry for 10 min. 8. Resuspend each sample in 50 μl TE buffer, and incubate for 3 to 4 hr at 55◦ C to allow the DNA to dissolve. 9. Store the DNA samples for at least 6 months at 4◦ C.

PCR IDENTIFICATION OF TARGETED hESC CLONES To identify targeted hESC clones, a PCR screening strategy is employed. This technique enables a large number of clones to be screened rapidly. The PCR screen is designed to amplify a novel junction fragment created by the correct homologous recombination event. One primer should anneal to sequences located in either the positive selection cassette or in the reporter gene (see Table 5B.1.1 for a list of recommended primers), while the second primer should prime from the target chromosomal sequences just beyond the homologous sequences used in the targeting vector (Fig. 5B.1.5). This method is also utilized to identify hESC clones in which the positive selection cassette has been excised from the genome (Basic Protocol 2), and to identify targeted lines following clonal isolation (Support Protocol 4). The robustness of the PCR amplification is in part related to the distance between the two primers and the composition of the DNA sequence being amplified. It is recommended that the amplified product be between 2 and 4 kb in length, and not contain long GC stretches. The addition of DMSO to the reaction may also assist in the amplification. Sequences of a similar size can be amplified from the wild-type locus (for example using primer ’b’ in Fig. 5B.1.5 and another primer binding to the wild-type locus ∼4 kb upstream) to assist in the optimization of the PCR reaction. The authors recommend using Platinum Taq DNA Polymerase High Fidelity (Invitrogen), and have found that this enzyme mixture regularly amplifies 1- to 5-kb sequences from genomic DNA using the reaction mixture listed in Table 5B.1.2.

SUPPORT PROTOCOL 3

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Table 5B.1.1 Primers Known to Anneal to Fluorescent Marker Genes and DrugResistance Cassettes

Gene

Sequences are 5 to 3

Annealing temperature

Direction of amplification

GFP

GTGCTGCTGCCCGACAACCACTAC CCGGTGAACAGCTCCTCGCCCTTGC

60◦ C 62◦ C

FWD (5 to 3 ) REV (3 to 5 )

RFP

CACAACACCGTGAAGCTGAAGGTGAC GTCACCTTCAGCTTCACGGTGTTGTG

65◦ C 65◦ C

FWD (5 to 3 ) REV (3 to 5 )

neo

CGATGCCTGCTTGCCGAATATCATG

60◦ C

FWD (5 to 3 )

hph

CTCCGCATTGGTCTTGACCAACTC

60◦ C

FWD (5 to 3 )

targeting vector

reporter

pos. select

exon 1

wild-type allele

exon 2

a targeted allele

reporter

b

pos. select

exon 2

Cre Recombinase c reporter

b exon 2

Figure 5B.1.5 A schematic representation of homologous recombination between the targeting vector and the wild-type allele. The wild-type allele represents a typical target gene containing two exons. The targeting vector contains sequences homologous to the genomic locus (orange lines and rectangles), as well as sequences encoding a reporter gene (green arrow), loxP sites (red triangles), and a positive selection cassette (blue rectangle). Homologous recombination between the wild-type allele and the targeting vector replaces a segment of exon 1 in the wild-type allele. A PCR assay is used to detect gene-targeted clones with primers (black arrows) corresponding to sequences within the positive selection cassette (a) and the endogenous wild-type allele (b). Only correctly targeted alleles will yield a PCR product. Following expression of Cre recombinase in the cells, the positive selection cassette is removed from the targeted allele. The loss of the positive selection cassette is confirmed by a second PCR using the same endogenous primer (b) and a primer corresponding to sequences within the reporter gene (c).

If any correctly targeted hESC clones are identified from the PCR screen, these cell lines should be maintained as colonies that are mechanically passaged, and stocks of each line frozen in liquid nitrogen. Genetically manipulated hESCs that are kept and grown as colonies retain their stem cell characteristics and are indistinguishable in appearance from the parental lines (Costa et al., 2005; Davis et al., 2008a).

Materials

Generation of hESC Reporter Knock-In Lines

PCR master mix (see Table 5B.1.2) containing: Autoclaved, distilled water 10× High Fidelity PCR Buffer (Invitrogen) 10 mM dNTP mixture (Sigma) 50 mM MgSO4

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Forward & Reverse Primers (see Table 5B.1.1) DMSO Platinum Taq High Fidelity DNA polymerase (Invitrogen) Genomic DNA from the hESC clones (Support Protocol 2 or the Alternate Protocol) hESC medium (see recipe) Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) 0.2-ml nuclease-free PCR tubes 1.5-ml nuclease-free microcentrifuge tubes Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Microcentrifuge DNA thermal cycler Gelatinized organ culture dishes Additional reagents and equipment for analyzing the amplification products by agarose gel electrophoresis (Voytas, 2001) and maintaining and expanding hESCs by both mechanical and enzymatic passaging (UNIT 1C.1) Set up and run the PCR 1. Organize the PCR tubes in the same layout as the DNA samples, on ice. 2. Determine the total number of DNA samples to screen. Include in this total both negative and positive control samples, if available. A possible negative control DNA sample is genomic DNA from nontransfected hESCs. Generally, a positive control DNA sample is not included. However, if a hESC line has previously been successfully targeted at that genomic locus, this can serve as a positive control.

3. Make a PCR master mix in a 1.5-ml nuclease-free microcentrifuge tube, on ice. A suggested cocktail is listed in Table 5B.1.2. Optimal concentrations for the PCR master mix will depend on the primer pair used and the DNA sequence being amplified, and must be determined empirically. This reagent assembly is best performed in a dedicated PCR preparation area using pipettors that have not been used to aliquot template DNA.

4. Aliquot 18 μl of the master mix into each PCR tube. Table 5B.1.2 PCR Master Mix

Amounta

Component

(n + 1) × 12.9 μl

dH2 O 10× High Fidelity PCR buffer 50 mM MgSO4

(n + 1) × 1.2 μl (n + 1) × 1 μl

DMSO 10 mM dNTPs 250 ng μl

(n + 1) × 2 μl

−1

forward primer

(n + 1) × 0.4 μl (n + 1) × 0.2 μl

250 ng μl−1 reverse primer

(n + 1) × 0.2 μl

5 U μl−1 Platinum Taq High Fidelity DNA polymerase

(n + 1) × 0.1 μl

Total

(n + 1) × 18 μl

a n equals the total number of reactions to be performed, including the positive and negative control

samples.

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5. To each PCR tube, add 2 μl of the appropriate genomic DNA sample. The DNA solutions are usually between 50 and 200 ng μl−1 and, in the 48-well plates, are often very viscous. These samples need to be mixed well with the pipet tip before removing 2 μl. To assist in tracking the PCR tubes into which the DNA samples have been added, leave the pipet tips containing the template DNA in the tube until all DNA samples have been aliquoted. Use a different pipettor to aliquot the DNA samples from the one used to aliquot the PCR master mix reagents.

6. Seal and briefly microcentrifuge the tubes 30 sec at 100 × g, room temperature. 7. Place tubes inside a thermal cycler, and enter cycle conditions. Commonly used amplification conditions are: 1 cycle:

3 min

30 to 40 cycles: 20 sec 30 sec 1 min (per kb of sequence being amplified) 1 cycle: 10 min

94◦ C 94◦ C 50◦ to 62◦ C 68◦ C

(initial denaturation) (denaturation) (annealing) (extension)

68◦ C

(final extension).

8. Visualize the PCR products by agarose gel electrophoresis (Voytas, 2001), and identify the hESC clones that have the desired genetic modification. If none of the hESC clones analyzed appear to be targeted, a second PCR should be performed using primers to amplify a similar-sized fragment in the same genomic region from the wild-type allele. A correct sized product obtained using this second set of primers indicates that it is unlikely that the absence of a PCR product from the screening PCR was a consequence of poor DNA template quality or low DNA concentration. If the primers specific to the wild-type allele fail to amplify the DNA fragment, further purification of the hESC DNA samples and/or optimization of the PCR protocol will be required prior to repeating the screening PCR.

Transfer the correctly targeted hESC clones onto organ culture dishes 9. Leave the hESC clones with the correct genetic modification to expand on the “Maintenance” 48-well plates, until the wells are ∼80% confluent (Fig. 5B.1.4B) or the colonies are beginning to differentiate. Change the hESC medium on the wells daily. Usually the hESC clones will be at this stage of growth by the time the PCR screening is complete.

10. One day before transferring the hESC clones, plate mitotically inactivated MEFs onto the center well of gelatinized organ culture dishes at a density of 6 × 104 per cm2 in 1 ml of MEF medium. Prepare enough organ culture dishes with MEFs, such that pieces from each targeted hESC clone can be plated onto two organ culture dishes.

11. On the day of transfer, replace the MEF medium on the organ culture dishes with 1 ml of hESC medium. 12. Mechanically fragment a hESC clone to be moved using the same technique described in step 12 of Support Protocol 1, and distribute the pieces of the clone across two organ culture dishes.

Generation of hESC Reporter Knock-In Lines

13. Repeat the above step with all genetically modified hESC clones that are to be transferred to organ culture dishes. Repeat the PCR screen on all positive clones that are identified to verify that the correct clone has been transferred from the maintenance plate.

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14. Maintain the reporter gene knock-in hESC lines as colonies that are mechanically passaged (UNIT 1C.1). Also expand the hESCs as a large-scale expansion by enzymatic passaging (UNIT 1C.1) for deletion of the positive selection cassette from the targeted lines (Basic Protocol 2). All targeted knock-in hESC lines generated at this stage should have stocks frozen in liquid nitrogen. A suggested method of cryopreservation is described elsewhere (Reubinoff et al., 2001).

REMOVING THE POSITIVE SELECTION CASSETTE FROM GENETICALLY MODIFIED hESCs BY TRANSIENT EXPRESSION OF Cre RECOMBINASE

BASIC PROTOCOL 2

Targeting vectors contain positive selection cassettes to facilitate the isolation of stably transfected cells. Following the isolation and identification of targeted lines, selection cassettes are removed because their continued presence may cause a number of undesirable effects. In genetically modified mice, the retention of the positive selection cassette can interfere with the expression of neighboring endogenous genes (Pham et al., 1996; Scacheri et al., 2001), while in reporter knock-in hESC lines it can result in the misexpression of the reporter (either by silencing or activating expression of the reporter; R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.). The removal of the positive selection cassette also offers the opportunity to subsequently retarget the remaining wild-type allele and to generate a homozygous knockout hESC line using the original targeting vector and drug selection strategy. If the positive selection cassette is flanked by loxP sites, expression of Cre recombinase in the genetically modified cells catalyzes the excision of the DNA sequence between the loxP sites (Sauer, 1993). This protocol describes a method for the transient transfection of a circular pEFBOS-creIRESpuro vector into hESCs using the lipofection reagent FuGENE 6 (Roche), and is an alternative to the transduction of a recombinant-modified Cre recombinase protein into the cells (Nolden et al., 2006). The transfected cells constitutively express both Cre recombinase and puromycin N-acetyltransferase and are selected for by the addition of the antibiotic puromycin to the hESC medium for 48 hr. This short selection period enriches for cells that have been transiently transfected with the pEFBOS-creIRESpuro vector and also allows sufficient time for Cre-mediated deletion of the antibiotic-resistance cassette to occur. Approximately 12 days after selection, the hESC colonies that have formed can be picked and expanded as described in Support Protocol 1. DNA is then extracted (Support Protocol 2) and the loss of the positive selection cassette is confirmed by PCR (Support Protocol 3).

Materials Gene-targeted hESCs containing a loxP-flanked positive selection cassette in gelatinized 75-cm2 tissue culture flasks between enzymatic passages 5 and 10 co-cultured with MEFs pre-seeded at a density of 1.5 × 106 /75-cm2 flask Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) hESC medium (see recipe) 60-mm gelatinized tissue culture dishes (see recipe) seeded with 3 × 104 /cm2 mitotically inactivated MEFs Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) FuGENE 6 Transfection Reagent (Roche) DMEM/F12 (Invitrogen) pEFBOS-CreIRESpuro expression vector (GenBank accession number EU693012; available on request from the authors’ laboratory; e-mail request to [email protected] or [email protected])

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Puromycin solution (Sigma), 10 mg/ml Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips 37◦ C incubator 15- and 50-ml sterile centrifuge tubes Refrigerated centrifuge 1.5-ml microcentrifuge tubes Additional reagents and equipment for performing a cell count (Phelan, 2006; UNIT 1C.3), transferring colonies to 48-well tissue culture plates (Support Protocol 1), extracting DNA from colonies (Support Protocol 2), and screening extracted DNA by PCR (Support Protocol 3) Seed feeders onto 60-mm dishes (day −2) 1. Supplement five 60-mm dishes with 2 ml gelatin to ensure that the gelatin coats the surface. 2. Allow to stand for 30 min at room temperature. 3. Aspirate gelatin and seed 6 × 105 feeders/dish in 4 ml MEF medium. 4. Store in a humidified incubator at 37◦ C, 5% CO2 until required.

Enzymatically passage genetically modified hESCs (day −1) 5. Harvest the genetically modified hESCs cultured in a 75-cm2 flask. Aspirate the hESC medium and rinse the flask with 5 ml CMF-PBS. Add 2 ml of trypsin or TrypLE Select to the flask and ensure that the dissociation solution coats the surface of the cells. Place the flask 4 hr at 37◦ C for 4 min and dislodge the hESCs from the flask with gentle tapping. 6. Add 8 ml of hESC medium to the flask and transfer the resuspended hESCs to a 15-ml centrifuge tube. 7. Pellet the cells by centrifuging the tube 3 min at 480 × g, 4◦ C, and remove the supernatant. 8. Resuspend the hESC pellet in 5 ml of fresh hESC medium and perform a cell count (Phelan, 2006; UNIT 1C.3). Subtract the number of MEFs (∼0.75 × 106 ) from the count to determine the total number of hESCs. Generally, a semi-confluent 75-cm2 flask will contain ∼4 × 106 hESCs, which is enough cells to seed five 60 mm-dishes with ∼0.8 × 106 hESCs per dish.

9. Transfer 4 × 106 hESCs to a 50-ml centrifuge tube. Add additional hESC medium so that the cells are resuspended in a total volume of 20 ml. 10. Distribute 4 ml of the hESC suspension into each of the five 60-mm dishes seeded the day before (steps 1 to 4) with MEFs at a density of 3 × 104 cells/cm2 . 11. Return the dishes to a humidified incubator at 37◦ C, 5% CO2 , and leave the hESCs to attach overnight.

Transfect the Cre recombinase expression vector into the hESCs (day 0) 12. Approximately 2 hr before performing the transfection, aspirate the medium on the 60-mm dishes and supplement the cells with 4 ml fresh hESC medium.

Generation of hESC Reporter Knock-In Lines

The dishes should be no more than 70% confluent with hESCs. If the cells are overconfluent, the hESCs may not transfect optimally with FuGENE. In addition, we have observed a reduction in efficacy of puromycin in eliminating untransfected hESCs if the hESCs are overconfluent at the time of antibiotic selection.

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13. Add 30 μl of FuGENE 6 Transfection Reagent directly to 470 μl DMEM/F12 in a sterile 1.5-ml microcentrifuge tube. Mix by flicking the microcentrifuge tube and incubate the complex for 5 min at room temperature The manufacturer advises that the undiluted FuGENE 6 Transfection Reagent should not come into contact with the walls of the microcentrifuge tube, as this can adversely affect the transfection efficiency.

14. Transfer 100 μl of the FuGENE 6/DMEM/F12 mixture into another microcentrifuge tube labeled “negative control.” This solution is applied to the fifth 60-mm dish that serves as a negative control.

15. Pipet 4 μl of a 1 μg/μl stock solution of pEFBOS-creIRESpuro plasmid DNA into the 400 μl FuGENE 6/DMEM/F12 mixture. Tap the microcentrifuge tube to mix the contents and leave at room temperature for a further 40 min. Incubate the 100 μl negative control centrifuge tube for the same period of time. The pEFBOS-creIRESpuro plasmid preparation must be pure and endotoxin-free. Transfection-grade plasmid preparation can be isolated using the Qiagen plasmid purification kits.

16. Into four of the hESC-containing 60-mm dishes, add 100 μl of the FuGENE/DNA complex mixture dropwise. Add the negative control into the fifth 60-mm dish. Swirl the dishes to ensure distribution over the entire surface, label all dishes appropriately, and return to the incubator. A negative control dish of hESCs for the FuGENE transfection is a good indicator of the kinetics and completeness of cell death in response to puromycin.

Select transiently transfected hESCs (day 1) 17. Supplement 50 ml of hESC medium with puromycin to a final concentration of 2 μg/ml. The optimal concentration of puromycin for selection may vary for different hESC lines. A titration should be performed to determine the minimum concentration required to eliminate untransfected hESCs within 48 hr of addition. For 2 days of selection, 50 ml of hESC medium containing puromycin should be sufficient stock. Discard any unused stock after 2 days.

18. Apply 5 ml of the selection medium to each of the five 60-mm dishes between 24 to 36 hr after FuGENE transfection. The hESCs should be ∼90% confluent.

19. Maintain puromycin selection for 48 hr and change the medium daily. Frequent medium changes are necessary during selection to remove the dead cells. After 48 hr, all the hESCs in the control dish should have died. If the dish still contains viable hESCs, either the concentration of puromycin was not sufficient to eliminate untransfected hESCs, or the dishes were too confluent with hESCs when selection was started.

20. Following puromycin selection, return to culturing the cells in hESC medium that does not contain puromycin. If the MEFs preseeded on the 60-mm dishes are puromycin-sensitive, they will need to be replaced once puromycin selection is stopped. Supplement each dish with ∼0.6 × 106 MEFs resuspended in hESC medium. If the MEFs are puromycin-resistant, only supplement with additional MEFs when gaps in the MEF layer appears. Try to maintain a MEF density of ∼2 × 104 viable MEFs/cm2 on the dishes.

21. Allow the colonies to grow for ∼12 days, changing the hESC medium daily.

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22. Once the colonies are ∼2-mm in diameter, transfer a minimum of 24 undifferentiated colonies to a 48-well tissue culture plate as described in Support Protocol 1 (steps 2 to 8). 23. Then extract DNA from these colonies (Support Protocol 2) and screen it by PCR (Support Protocol 3) to confirm that the positive selection cassette has been removed from the hESCs. This is achieved by choosing a 5 primer that anneals to sequences within the reporter gene, and a 3 primer to sequences 3 of the antibiotic-selection cassette (Fig. 5B.1.5). A genomic DNA sample from targeted hESCs that still contain the selection cassette can be included in the screen to visualize the difference in size between PCR products still containing the positive selection cassette and those in which the cassette has been excised. Additional PCR screens using primers specific to the Cre recombinase expression vector to verify that the plasmid did not integrate into the genome of the resulting colonies, and a PCR using primers specific to the antibiotic-resistance cassette to exclude the presence of residual unexcised cells, should be performed. The sensitivity of the hESCs to geneticin and puromycin can also be confirmed later by re-exposing an aliquot of the cells to the selection agents. Two or three of these reporter knock-in hESC clones that have had the positive selection cassette removed should be returned to organ culture dishes for maintenance and expansion. These lines should also be cloned as described in Support Protocol 4 to ensure that the targeted hESC lines used in future applications contain a targeted locus in which the antibiotic selection cassette has been excised. SUPPORT PROTOCOL 4

CLONAL ISOLATION OF hESCs BY SINGLE-CELL DEPOSITION FLOW CYTOMETRY The procedures for generating a targeted hESC line and removing the positive selection cassette support the clonal growth of hESCs. However, they do not necessarily exclude the possibility that the resulting hESC colonies arose from more than one cell. This protocol uses flow cytometry to derive single-cell clones from the existing parental hESC lines. Despite the low cloning efficiency of hESCs reported in the literature (Sidhu and Tuch, 2006), we routinely achieve a single-cell cloning frequency of 4% to 8% using hESCs adapted to enzymatic passage as we have described in UNIT 1C.1. Viable hESCs are selected by size gating and exclusion of the dye, propidium iodide (PI), and single cells are deposited directly into individual wells of 96-well plates using a flow cytometer. Correct gene targeting and absence of the positive selection cassette is confirmed in the resulting clones by PCR. It is possible to merge this protocol with the procedure to remove the positive selection cassette (Basic Protocol 2), reducing the time required to generate the final targeted hESC line. A description of this integrated procedure is provided elsewhere (Davis et al., 2008b). Subcloning ensures that the resulting targeted hESC lines consist of cell populations with identical genetic constitutions. This method is also a useful approach for ensuring a homogeneous diploid cell population, and selecting hESC lines with a uniform, undifferentiated morphology.

Materials

Generation of hESC Reporter Knock-In Lines

0.1% (w/v) gelatin solution (see recipe) Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) Genetically modified hESCs generated from Basic Protocol 2 in 75-cm2 tissue culture flasks between enzymatic passages 5 and 10

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75-cm2 gelatinized tissue culture flasks (see recipe) seeded with mitotically inactivated MEFs at 1 × 104 /cm2 for passaging hESCs prior to cloning hESC medium (see recipe) Recombinant human FGF2 (see recipe) Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) Propidium iodide (PI) solution (see recipe) Flat-bottomed 48-well tissue culture plates, gelatinized and seeded with mitotically inactivated MEFs at 0.75 × 106 /plate Liquid nitrogen 96-well flat-bottom tissue culture-treated plates and lids 37◦ C, 5% CO2 incubator 15-ml sterile centrifuge tubes Refrigerated centrifuge 5-ml sterile round-bottom polystyrene FACS tubes (12 × 75–mm) with 35-μm cell-strainer caps and with snap lids (Falcon) Parafilm M (Pechiney Plastic Packaging) or equivalent Flow cytometer with single-cell deposition function, e.g., FACSVantageSE-DiVa system (Becton Dickinson) or equivalent Inverted microscope Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Stereomicroscope Additional reagents and equipment for propagating hESCs in bulk culture (UNIT 1D.3), extracting DNA from colonies (Support Protocol 2), screening extracted DNA by PCR (Support Protocol 3), maintaining and expanding the genetically modified lines (UNIT 1C.1), and for cryopreserving hESCs (Reubinoff et al., 2001) Feeder reduce the hESCs to be subcloned and prepare the 96-well plates (day −1) 1. Add ∼50 μl of 0.1% gelatin solution to each well on ten 96-well plates. Leave the plates at room temperature for 15 min. 2. Aspirate the gelatin solution from the wells. 3. Resuspend MEFs in MEF medium such that the final concentration is 2 × 105 MEFs/ml. Aliquot 50 μl of the MEF-containing medium into each well. The final density of MEFs is ∼1 × 104 MEFs/well.

4. Passage hESCs for subcloning into a 75-cm2 flask containing MEFs seeded at a density of 1 × 104 /cm2 (UNITS 1C.1 & 1D.3). Harvest the hESCs as described in steps 1 to 3 of Basic Protocol 2. Resuspend the pelleted hESCs in hESC medium and transfer three-quarters of the cell suspension to the 75-cm2 flask. To maintain the hESC line, the remaining cells can be plated into another tissue culture flask containing MEFs at the normal density (2 × 104 MEFs/cm2 ). The passaging of the hESCs onto low-density MEFs increases the hESC:MEF ratio, improving the number of hESC subclones obtained following single-cell deposition.

5. Incubate both the 96-well plates and the flasks containing the passaged hESCs in a humidified incubator at 37◦ C, 5% CO2 overnight.

Subcloning of the gene-targeted hESC line (day 0) 6. Supplement 300 ml of hESC medium with additional FGF2 so that the final concentration of FGF2 is 40 ng/ml.

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The higher concentration of FGF2 supports the growth and expansion of the hESCs, and also suppresses the spontaneous differentiation that can occur during the early establishment of the single-cell clones.

7. Aspirate the MEF medium from the wells of the multi-well plates, and aliquot 100 μl of hESC medium containing 40 ng/ml FGF2 into each well. Return the plates to the incubator until required for use with the flow cytometer.

Harvest hESCs 8. Harvest the hESCs to be subcloned. Briefly, remove the medium from the 75-cm2 flask, rinse the cells with 5 ml CMF-PBS, add 3 ml of the dissociating agent (either TrypLE Select or trypsin) and return the flask to the 37◦ C incubator for 4 to 5 min. 9. Gently tap the base of the flask to dislodge the hESCs and add 10 ml hESC medium (without the additional FGF2) to the flask. Mix and transfer this cell suspension to a 15-ml centrifuge tube. 10. Pellet the cells by centrifuging the tube for 3 min at 480 × g, 4◦ C. 11. Remove the supernatant and resuspend the hESCs in 1 ml of hESC medium (without the additional FGF2). 12. Filter the hESCs by passing the cell suspension through a 35-μm cell-strainer cap attached to a sterile FACS tube. To help the dissociated hESCs pass through the cell strainer, cover the caps with a small square of Parafilm and centrifuge the tube for 3 min at 480 × g, 4◦ C, to pellet the cells. This step removes cell clumps and cellular debris and ensures that the hESCs form a single-cell suspension. Covering the caps the Parafilm helps to keep the cells sterile.

13. Return the tube to the tissue culture hood and discard the Parafilm and cell strainer cap. 14. Carefully aspirate the supernatant and resuspend the hESCs in 1.5 ml of hESC medium supplemented with 40 ng/ml FGF2. To this mixture, also add 15 μl of 100 μg/ml propidium iodide (PI) solution. Flick the tube to mix. 15. Recap the FACS tube with a sterile FACS tube cap and store at 4◦ C or on ice until required. Sealing the FACS tube prevents desiccation and keeps the cell suspension sterile.

16. Identify the viable hESCs by size gating and exclusion of PI on a flow cytometer whose lines have been sterilized. Deposit single cells directly into each well of the ten 96-well plates. Although not routinely used in our laboratory, the cloning efficiency of the hESCs can be further improved by treating the cells with the Rho-associated kinase inhibitor (ROCKi), Y-27632 (Watanabe et al., 2007). We have had success by adding the inhibitor at 2 μM to the hESC medium in the 96-well plates at the time of sorting.

17. Return the plates to a humidified incubator at 37◦ C, 5% CO2 . 18. Four days after seeding the plates with hESCs, supplement the medium in each well with an additional 50 μl of hESC medium containing 40 ng/ml FGF2. We do not advocate the continued inclusion of ROCKi in the culture medium.

Generation of hESC Reporter Knock-In Lines

Identify wells containing hESC colonies (day 8) 19. Use an inverted microscope to identify wells containing viable hESC colonies. Carefully aspirate the medium from these wells and replace with 100 μl of hESC

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medium containing 40 ng/ml FGF2, as well as fresh MEFs at a concentration of 3 × 104 MEFs/ml. 20. Allow the colonies to continue growing. Replace the hESC medium on the wells containing viable hESC colonies daily. When the colonies are ∼2-mm in diameter, the hESCs can be transferred to and expanded on 48-well tissue culture plates as described in step 21. However, if the colonies begin to differentiate before they reach that size, they should be transferred sooner.

Transfer the hESC colonies onto 48-well plates (between days 16 to 18) 21. Determine the total number of viable hESC colonies on the 96-well plates. On the day before moving the colonies, seed enough 48-well plates with ∼0.75 × 106 MEFs/gelatinized plate (∼2 × 104 /cm2 ) so that each hESC colony can be transferred to an individual well. Incubate the 48-well plates overnight at 37◦ C, 5% CO2 to allow the MEFs to attach. Generally, we observe between 40 to 80 colonies spread over the ten 96-well plates.

22. Replace the medium on the 48-well plates with 200 μl of hESC medium supplemented with 10 ng/ml FGF2 per well. The concentration of FGF2 in the hESC medium can be returned to 10 ng/ml from this stage onwards.

23. Fragment a hESC colony into multiple small cell clumps with a 200-μl plugged tip attached to a pipettor. Transfer these pieces into a well on one of the 48-well plates. This can be performed macroscopically and the well on the 96-well plate checked afterwards under a stereomicroscope to confirm that the entire hESC colony detached and was transferred.

24. Discard the pipet tip after harvesting each colony. 25. Repeat steps 23 and 24 to transfer the remaining hESC colonies growing on the 96-well plates. The 96-well plates can be discarded once all the viable hESC colonies have been transferred.

Maintain and expand the clones 26. Maintain replicate and expand the hESC colonies as described in steps 8 to 17 in Support Protocol 1. 27. Extract DNA from these colonies (Support Protocol 2) and perform a PCR screen (Support Protocol 3) to confirm the subclones are correctly targeted, have the positive selection cassette excised from the genome, and did not integrate Cre recombinase into the genome (Basic Protocol 2, step 21, annotation). Discard any subclones that are negative for this screen. 28. From the remaining subclones, choose two or three that are growing well and appear undifferentiated in culture, and return these colonies to organ culture dishes. 29. Maintain and expand these genetically modified lines using the protocols described in UNIT 1C.1. 30. Confirm that these subclones have maintained a stem cell phenotype and a normal karyotype. 31. Cryopreserve stocks of these lines (Reubinoff et al., 2001). Generic Manipulation of Stem Cells

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REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

DNA lysis buffer 100 mM Tris·Cl, pH 8.0 200 mM NaCl 5 mM EDTA, pH 8.0 Add 0.2% (w/v) SDS powder (Sigma) last At time of use, dispense the required volume and add 200 μg/ml proteinase K (see recipe) to the solution Store the solution without proteinase K at room temperature indefinitely Gelatin, 0.1% (w/v) Add 0.5 g of gelatin powder (from porcine skin; Sigma) to 500 ml distilled water and autoclave to dissolve and sterilize. Store up to 6 months at room temperature.

Gelatinization of plates and flasks Prior to addition of MEFs, add enough 0.1% (w/v) gelatin solution (see recipe) to cover the base of all plates/flasks. Let stand for 10 min at 37◦ C to coat the surface and remove by aspiration immediately prior to addition of MEFs.

hESC medium DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (Invitrogen) 10 mM non-essential amino acids (Invitrogen) 2 mM L-glutamine or GlutaMaxI (Invitrogen) 1× penicillin/streptomycin (200× stock; Invitrogen) 100 μM 2-mercaptoethanol 10 ng/ml FGF2 (see recipe) Filter sterilize using a 0.22-μm Stericup filtration unit (Millipore) Store up to 1 week at 4◦ C MEF medium DMEM (4.5 g/liter glucose, without L-glutamine and sodium pyruvate; Invitrogen) containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine (Invitrogen) 1× penicillin/streptomycin (200× stock; Invitrogen) Filter sterilize and store up to 4 weeks at 4◦ C Propidium iodide (PI) Dissolve PI (Sigma) in CMF-PBS to a final stock concentration of 100 μg/ml (100× stock). Filter sterilize and store for at least 12 months at 4◦ C.

Proteinase K Dissolve proteinase K powder (Sigma) in H2 O to a final stock concentration of 20 μg/μl (20 mg/ml; 100×). Dispense into 1-ml aliquots and store for at least 12 months at –20◦ C. Generation of hESC Reporter Knock-In Lines

Recombinant human basic fibroblast growth factor (FGF2) Reconstitute lyophilized rhFGF2 (PeproTech) to a final stock concentration of 10 μg/ml in CMF-PBS. Store up to 6 months at –80◦ C.

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Soybean trypsin inhibitor Dissolve the soybean trypsin inhibitor stock powder (Sigma) in CMF-PBS to a final concentration of 1 mg/ml. Filter sterilize and dispense into 5- or 10-ml aliquots and store up to 12 months at −20◦ C. Once thawed, the solution can be stored for 1 to 2 weeks at 4◦ C.

Tris/EDTA (TE) buffer, pH 8.0 10 mM Tris·Cl, pH 8.0 1 mM EDTA, pH 8.0 Store the solution at room temperature indefinitely Trypsin, 0.125% (w/v) Supplement Trypsin/EDTA [0.25% (w/v) trypsin EDTA. 4Na, Invitrogen] with 2% (v/v) chicken serum (Hunter antisera). Decant into 5-ml aliquots and store up to 12 months at −20◦ C. To use, thaw an aliquot and add an equal volume of CMF-PBS. Once thawed, the solution can be stored up to 4 weeks at 4◦ C.

COMMENTARY Background information Human embryonic stem cells are derived from the inner cell mass of the pre-implantation blastocyst stage embryo (Thomson et al., 1998; Reubinoff et al., 2000). These cells exhibit two key characteristics. Firstly, they can be maintained and expanded in vitro for extended periods of time as undifferentiated cells while preserving their original karyotype. Secondly, they are pluripotent, and therefore have the capacity to differentiate into various cell types representing the three germ layers, both in vivo and in vitro (Amit et al., 2000). The ability to transform hESCs into multiple lineages in culture provides opportunities to examine human embryonic development in vitro, generate specific cells and tissues for therapies or drug screening, and identify chemical compounds that influence a specific developmental process. The identification of a cohort of lineagespecific markers assists in the directed differentiation of hESCs towards specific cell types. When appropriate antibodies for specific cellsurface markers are available, fluorescenceactivated cell sorting (FACS) may be used to purify viable hESC derivatives. Where antibodies for suitable cell surface markers are unavailable, or the lineage-specific markers are intracellular, reporter genes can be targeted to loci whose expression marks critical developmental milestones, facilitating the isolation of viable cell populations that would otherwise be inaccessible. These purified subpopulations can then be further differentiated or expanded in vitro, or transplanted and tracked

in vivo. These approaches have contributed significantly to our understanding of lineage specification in differentiating mouse ESCs (Fehling et al., 2003; Ying et al., 2003; Ng et al., 2005; Micallef et al., 2005, 2007; Gadue et al., 2006). Until recently, similar strategies have been unachievable with hESCs, partly due to suboptimal culturing conditions leading to poor transfection and single-cell cloning efficiencies. However, several studies have now described the introduction of selectable markers into hESCs using a variety of different approaches (Giudice and Trounson, 2008, and references therein). These include using vectors or viruses to randomly integrate reporter genes regulated by lineage-specific promoter fragments into hESCs. While these methods can introduce new genetic material into the genome at relatively high frequencies, there are several caveats associated with random integration, the foremost being that expression of the reporter may not faithfully reflect expression of the endogenous gene. Other concerns include the possibility of disrupting normal gene functions, and that elements other than the promoter fragments included in the construct may be required for accurate expression of the reporter. Furthermore, unlike mouse ESC lines, in which transgenic lines can be validated by examining reporter gene expression patterns in chimeric mice, validation of analogous hESC lines relies on examination of reporter expression in vitro or the more limited in vivo setting afforded by xenogenic transplants.

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Gene targeting utilizes the cellular DNA repair machinery to integrate transgenes, such as reporter genes, by homologous recombination into a specific site in the genome. The reporter gene is regulated in the same manner as the endogenous gene from the remaining wild-type allele and is thus expressed at the same time and in the same cells. Therefore, although homologous recombination is technically challenging, it produces expression patterns that more accurately reflect native gene expression and is preferable to random integration for the production of reporter cell lines. While electroporation is the chosen method for gene targeting in mouse ESCs, initially it was considered that this technique was unlikely to be successful in hESCs due to the resulting high mortality (Eiges et al., 2001). Modifications to both the culturing of hESCs and to the standard mouse ESC electroporation protocol have formed the basis of a successful targeting method (Costa et al., 2007). While it is possible to obtain homologous recombinants in hESCs using cationic reagents, this occurs at a very low frequency (10−8 ; Urbach et al., 2004). Using the electroporation procedure described in this unit, the stable transfection frequency is consistently between 2 × 10−6 and 5 × 10−5 (i.e., 20 to 500 colonies per 107 transfected cells), of which 1.3% to 14.6% of the stable transfectants are correctly targeted, depending on the locus and the specific hESC line (Costa et al., 2007). While the stable transfection frequency in hESCs is around 100-fold lower than what is usually observed in mESCs (Vasquez et al., 2001), the frequency of homologous recombination within stable transfectants appears comparable between the two species. During the differentiation of hESCs, a panel of markers are generally used to screen and identify lineage-specific cell types. Engineering dual or multiple reporter knock-in hESC lines would enable the identification of pools of cells that share common markers and improve our understanding of the molecular and cellular mechanisms that govern lineage specification. Results from these lines of analyses can subsequently be utilized to differentiate unmodified hESC lines to cell types with potential clinical applications.

Critical Parameters and Troubleshooting Generation of hESC Reporter Knock-In Lines

Gene targeting strategy The generation of reporter knock-in hESC lines is a challenging task, as illustrated by the

limited number of reports describing successful gene targeting in hESCs. In our laboratory, we aim to generate several independent genetargeted clones in more than one hESC line in order to demonstrate the generalizability of the experimental findings. The planning of the gene targeting strategy should consider both vector design and the processes involved in the characterization and validation of the newly formed line. The Strategic Planning section contains guidelines regarding the structure of the targeting vector. One parameter that appears to influence the frequency of homologous recombination in hESCs is the length of the homology arms (Zwaka and Thomson, 2003). Having at least one homology arm >6 kb improves the targeting frequency (Zwaka and Thomson, 2003; A.G. Elefanty and E.S. Stanley, unpub. observ.). However, the origin of the homology arms does not appear to significantly influence the targeting frequency between different hESC lines (Costa et al., 2007). The reduced requirement for the homology arms to be derived from isogenic DNA may reflect the lower frequency of polymorphisms that we have observed between different human DNA isolates compared to DNA from different inbred mouse strains (R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.). Therefore, we find that commercially available bacterial artificial chromosome DNA is a convenient source of genomic DNA. Similar to mouse ESCs, the frequency of homologous recombination in hESCs also appears to be locus dependent (Hasty et al., 1994; Costa et al., 2007). If homologous recombination is not obtained after the initial electroporation, the experiment should be repeated until at least 500 stably transfected clones have been screened. If targeting is still unsuccessful, we have achieved success by altering the length of the homology arms. Another alternative is to target a different region of the gene. If the generation of a targeted line in which the loss of one allele is anticipated to result in haplotype insufficiency, targeting the 3 untranslated region of the gene with a vector including an Internal Ribosomal Entry Site (IRES) upstream of the reporter coding sequences may circumvent such a problem. A fluorescent reporter marker, usually a green fluorescent (GFP) or red fluorescent (RFP) protein, is the typical reporter gene used to generate a reporter knock-in hESC line (Zwaka and Thomson, 2003; Irion et al., 2007; Davis et al., 2008a). The fluorescent protein provides an easy visual identifier of cells

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corresponding to the desired population, as well as enabling the quick assessment of how the differentiating cells respond to different stimuli. In hESCs, the expression of GFP can persist beyond the timeframe that the lineagespecific marker is detected, making it also useful for lineage tracing experiments (Davis et al., 2008a). In instances where this function is not desirable, another reporter such as nonfunctional versions of cell surface receptors (e.g., CD4 or CD25) could be used, as has been done in mESCs (Yasunaga et al., 2005; Gadue et al., 2006). While cells expressing surface markers cannot be readily visualized, antibodies are available that enable the detection and isolation of these cells by immunofluorescence or flow cytometry. Alternatively, if direct visualization of the reporter knock-in cells is a requirement, then an unstable form of GFP that has a significantly reduced half-life could be utilized (Corish and Tyler-Smith, 1999). However, the cumulative fluorescence from such a GFP may also be reduced, making direct visualization of expressing cells difficult. Culturing of hESCs A homogeneous population of undifferentiated hESCs is required for these protocols. Most culturing systems consist of hESCs grown on mitotically inactivated MEFs, which provide some of the necessary factors for the maintenance and survival of hESCs in vitro. The quality of the mitotically inactivated MEFs can therefore significantly affect the hESCs culture. The hESCs should be routinely analyzed by flow cytometry for the expression of a panel of stem cell markers. Experience in passaging and maintaining undifferentiated hESCs in bulk culture is recommended before embarking on the protocols described in this unit. It is vital that the hESCs are adapted to enzymatic passaging as single cells before being transfected or cloned. Care should always be taken when enzymatically harvesting the hESCs. The dissociation treatment should last only as long as necessary to dislodge the cells from the tissue culture plastic ware. The clumps can then be gently broken up by trituration into a suspension of (predominantly) single cells. Prolonged enzymatic passaging of hESCs can select for cells adapted to this culturing technique and even result in the acquisition of chromosomal aberrations that offer a selective advantage (Draper et al., 2004). When generating genetically modified lines, the parental cell lines should be enzy-

matically passaged no more than 10 times before use. The hESCs are always passaged the day prior to an application so that the following day the cultures are semi-confluent and therefore actively proliferating before being utilized. The cells are also supplemented with fresh medium a few hours before transfecting or cloning the cells. Following the generation or cloning of gene-targeted hESC lines, the cells should be returned to organ culture dishes and maintained as dense colonies that are mechanically passaged once a week. Re-entering the organ culture phase reduces the chance of obtaining chromosomally abnormal lines. Stocks of the lines should also be frozen in liquid nitrogen. Electroporation of the targeting construct and selection of stably transfected hESCs Viable hESC colonies should emerge in culture within 2 days following electroporation (Fig. 5B.1.3B). Very high mortality levels observed following electroporation might be a consequence of excessive or rough handling of the cells throughout the protocol. Also, ensure the hESCs are resuspended in ice-cold PBS prior to electroporation and later transferred to prewarmed hESC medium (37◦ C). The electroporated cells should be washed with hESC medium before plating to remove any cellular debris that might be detrimental to the surviving hESCs. We believe that these steps improve the recovery level of the hESCs following electroporation. High concentrations of plasmid DNA can also adversely affect hESC viability and using a reduced quantity of DNA can lower mortality during electroporation, particularly if the cell count is below 1 × 107 . An endotoxin-free plasmid purification kit should be used when preparing the plasmid DNA. Rapid differentiation of the hESCs after electroporation suggests that the culture conditions may have been suboptimal or the electroporation parameters were too harsh. Under these circumstances, often the cells cannot be rescued and it is best to discard the cells and re-commence the electroporation procedure. To minimize the risk of differentiation in future transfections, the MEF density on the dishes and/or the concentration of FGF2 in the hESC medium could be increased. In addition, the hESC medium should be changed more frequently, and the dishes supplemented with MEFs earlier on during the drug selection period.

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Following selection, drug-resistant colonies should be visible within 1 week. If no colonies have formed, then the positive selection cassette in the targeting construct may be nonfunctional. Prior to electroporation, the expression of the positive selection cassette should be confirmed in another cell type, such as the human embryonic kidney (HEK) 293T line. Another possibility for the absence of hESC colonies is a failure of the electroporation procedure to transfect the vector into the cells. An abnormally short time constant (e.g., <10 msec when cells are transfected at 250 V and 500 μF) and/or absence of a flocculent precipitate of dead cells immediately after the electroporation can indicate this. A parallel electroporation into the same hESC line using a vector (such as pEFBOS-GFPneo or human β-actin GFP-IRES-Neo) that has been demonstrated to work in hESCs can be used as a positive control (Costa et al., 2005, 2007). If the hESCs fail to die during drug selection, it is likely that the drug concentration is too low and should be increased. Always perform a titration of the drug on the parental hESC line to ensure the optimal concentration is used for selection.

Generation of hESC Reporter Knock-In Lines

PCR screening to identify hESCs that have undergone homologous recombination A PCR-based method for detecting homologous recombinants is the most efficient method for screening large numbers of clones, and is very reliable. However, the choice of PCR primers is one of the most crucial aspects in achieving the desired sensitivity of the PCR screen. The primers should be ∼24 bp in length, with the GC content between 50% and 60%. The GC residues should be equally distributed throughout the primer, and there should be no obvious secondary structures formed. The presence of either a G or C residue at the 3 end of the primers provides a more stable extension start site for the Taq DNA polymerase. The PCR screening strategy also requires optimization for each primer pair and locus. One approach is to design a primer against a region of the endogenous genomic sequence that is replaced by the targeting vector during the recombination event, and use this primer in combination with the endogenous screening primer. The best PCR conditions are then determined based on amplifying a similarsized DNA fragment from the same loci where targeting will occur, but from the wild-type allele.

Alternatively, a PCR optimization strategy based on a “mock knockout construct” can be performed (Kontgen and Stewart, 1993). Genomic DNA from ∼1 × 106 hESCs is mixed with ∼1 μg linearized targeting vector and is used as the substrate to determine the best PCR screening conditions for homologous recombinants. The inclusion of genomic DNA samples from the parental hESC lines in the PCR screen can serve as useful controls for false positives. Removal of positive selection cassette Once correctly targeted hESC clones are obtained and confirmed to be karyotypically normal, the positive selection marker is then removed from the genomic locus to prevent it from interfering with the expression of the reporter and neighboring genes. If the selection cassette is flanked by loxP sequences, then transient transfection of the pEFBOScreIRESpuro expression vector will result in the excision of all sequences sandwiched between the loxP sequences. The drug puromycin is added to the culture medium to select for hESCs that are expressing the transfected vector. After 2 days of selection, small clusters of live hESCs should be visible by microscopy. If no groups of hESCs are visible, steps should be undertaken to confirm that the vector has transfected into the cells. Co-transfecting a vector carrying a constitutively expressed fluorescent reporter can optimize the transfection protocol. A titration to determine the optimal concentration of puromycin required for selection should also be performed. A negative control consisting of a dish of untransfected hESCs should always be included. These cells should all die within 48 hr of puromycin exposure. If the dishes containing the hESCs transfected with the pEFBOScreIRESpuro expression vector remain confluent following 2 days of puromycin selection, it is likely that the initial seeding density of the cells was too high for effective selection. The vector should not be transfected into cultures of hESCs that are >70% confluent or are not growing as a monolayer of cells. Because of the short exposure time to puromycin, this selection method can be used even if the supporting MEF cell line is puromycin sensitive. However, the dishes must be replenished with fresh MEFs immediately post puromycin selection to support the growth of the remaining viable hESCs. Again, the initial procedure to identify the hESC clones that have excised the positive

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selection marker from the targeted locus relies on PCR-based screening. Using a primer against sequences in the reporter gene in combination with a primer that matches sequences 3 of the positive selection marker, a PCR product should be amplified regardless of whether the cassette has been excised or not. If no product is obtained, this suggests that either the PCR conditions were suboptimal, or the original gene-targeted hESC line was not clonal and included nontargeted hESCs. To optimize the PCR conditions, include as a positive control a sample of DNA from the gene-targeted hESC line that still contains the positive selection marker. If the original gene-targeted hESC line is not clonal, the researcher should continue to screen colonies that were puromycin-resistant by PCR until they identify clones that are correctly targeted and have lost the positive selection cassette. These hESCs will also need to be clonally isolated using single-cell deposition flow cytometry. PCR should be performed to confirm that the Cre recombinase expression vector did not integrate into the genome. It is also recommended that the chosen hESC clones be reexposed to the selection agents to confirm their sensitivity. Generally, almost all of the colonies screened have the positive selection cassette excised (Davis et al., 2008b). Cloning It is important that all reporter knock-in hESC lines that will be used as reagents in future experiments are clonally isolated. This ensures that the final cell population is homogeneous with respect to the targeted locus and the excision of the antibiotic-selection cassette. For example, a mixed population of targeted and nontargeted hESCs could result in the misclassification of cell types if identification is based on the expression of the reporter gene. The cloning efficiency of hESCs using single-cell deposition flow cytometry varies between cell lines. Always increase the concentration of FGF2 in the hESC medium to 40 ng/ml during cloning. This helps to grow and expand colonies from individual hESCs and reduces the level of spontaneous differentiation (Amit et al., 2000; Xu et al., 2005). If the colonies still spontaneously differentiate on the 96-well plates, increase the frequency of hESC medium changes and/or the density of the mitotically inactivated MEF cells in the wells.

If less than 10 viable hESC colonies are obtained following the isolation of ∼1000 individual hESCs, 2 μM of the Rho-associated kinase inhibitor, Y-27632, could be added to the cells following single-cell deposition. This could improve the level of survival of the hESCs; however, the poor cloning efficiency could also be due to the ratio of MEFs to hESCs being too high in the sorted population. Reducing the concentration of feeder cells by passaging the hESCs onto a lower density of MEFs (∼1 × 104 /cm2 ) the day before singlecell cloning reduces the background feeder count. On the day of single-cell deposition, the flasks should be ∼80% confluent with hESCs.

Anticipated Results In most cases, if the guidelines and protocols described in this unit are adhered to, a reporter knock-in hESC line will be generated in which the expression of the reporter accurately reflects that of the targeted gene. The hESCs will also retain their stem cell characteristics and remain karyotypically normal. At the time of writing, the authors’ laboratory has used these protocols to generate reporter knock-in hESC lines at nine different loci, with multiple targeted clones in two independent hESC lines for most cases. The electroporation protocol (Basic Protocol 1) should yield stably transfected hESC clones at a frequency of 20 to 500 per 107 input hESCs. However, the frequency of homologous recombination amongst these surviving clones will vary depending on the vector used and the genomic locus being targeted. In the authors’ laboratory, this frequency has ranged from 1.3% to 14.6%. The method for the removal of loxPflanked positive selection cassettes from the genome of genetically modified hESC lines (Basic Protocol 2) has proven to be very successful. On average, 62 puromycin-resistant colonies are obtained per 1 × 106 hESCs transfected with the pEFBOS-creIRESpuro expression vector, with >95% of the resulting colonies screened having the selection cassette removed. The cloning efficiency of the hESCs using single-cell deposition flow cytometry is generally between 4% and 8%.

Time Considerations Once the targeting vector has been constructed, it will take a minimum of 6 months to generate and validate a reporter knock-in hESC line. However, it could take several more

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Generation of hESC Reporter Knock-In Lines

months if a correctly targeted hESC clone is not obtained from the first electroporation attempt. On average, it takes 1 year from designing and constructing the targeting vector to having an established and proven targeted reporter knock-in hESC line. The electroporation and subsequent selection of hESC clones that have integrated the targeting construct takes ∼2 1/2 weeks. The expansion of these clones and isolation of DNA for PCR screening requires a further 2 weeks of culturing. This can be reduced by ∼1 week if DNA is prepared directly from the hESC clones on the “Primary” 48-well plates, prior to replicating plates. The PCR screen should take no more than 2 days. Because this entire procedure takes ∼1 month, it is recommended that several consecutive electroporations be initiated at weekly intervals, even before the PCR screening results from the first transfection are known. Once a correctly targeted hESC clone is obtained, it will take at least 4 weeks of culturing to expand the clone for removal of the positive selection cassette. Additionally, these hESC lines should be cryopreserved and karyotyped, and homologous recombination confirmed by Southern blot analysis. The procedure for the removal of the positive selection cassette and subsequent confirmation by PCR will also take ∼1 month. The transfection of the pEFBOS-creIRESpuro vector and selection by puromycin requires 5 days. The transfected hESCs then require at least another 10 days before they can be transferred to 48-well plates for expansion. After the expansion of these clones (∼2 weeks), the extraction of DNA and PCR analysis will take 3 days. The expansion of two or three hESC clones in which the selection cassette has been removed will take another month. The cloning of these lines by single-cell deposition flow cytometry and confirmation that the cells contain the correct genetic modification requires a further 4 weeks. If the cloning protocol is integrated into the procedure for removing the positive selection cassette, it is possible to reduce the length of time required to generate a reporter knock-in hESC line by ∼2 months.

Council (NHMRC) of Australia. AGE is a Senior Research Fellow of the NHMRC.

Acknowledgements

Eiges, R., Schuldiner, M., Drukker, M., Yanuka, O., Itskovitz-Eldor, J., and Benvenisty, N. 2001. Establishment of human embryonic stem celltransfected clones carrying a marker for undifferentiated cells. Curr. Biol. 11:514-518.

We thank Robyn Mayberry, Kathy Koutsis, and Amana Bruce for the provision of hESCs. This work was supported by the Australian Stem Cell Centre (ASCC), the Juvenile Diabetes Research Foundation (JDRF), and the National Health and Medical Research

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Current Protocols in Stem Cell Biology

Fehling, H.J., Lacaud, G., Kubo, A., Kennedy, M., Robertson, S., Keller, G., and Kouskoff, V. 2003. Tracking mesoderm induction and its specification to the hemangioblast during embryonic stem cell differentiation. Development 130:4217-4227. Gadue, P., Huber, T.L., Paddison, P.J., and Keller, G.M. 2006. Wnt and TGF-beta signaling are required for the induction of an in vitro model of primitive streak formation using embryonic stem cells. Proc. Natl. Acad. Sci. U.S.A. 103:16806-16811. Giudice, A. and Trounson, A. 2008. Genetic modification of human embryonic stem cells for derivation of target cells. Cell Stem Cell 2:422-433. Gu, H., Zou, Y.R., and Rajewsky, K. 1993. Independent control of immunoglobulin switch recombination at individual switch regions evidenced through Cre-loxP-mediated gene targeting. Cell 73:1155-1164. Hasty, P., Crist, M., Grompe, M., and Bradley, A. 1994. Efficiency of insertion versus replacement vector targeting varies at different chromosomal loci. Mol. Cell Biol. 14:8385-8390. Hug, B.A., Wesselschmidt, R.L., Fiering, S., Bender, M.A., Epner, E., Groudine, M., and Ley, T.J. 1996. Analysis of mice containing a targeted deletion of beta-globin locus control region 5 hypersensitive site 3. Mol. Cell Biol. 16:29062912. Irion, S., Luche, H., Gadue, P., Fehling, H.J., Kennedy, M., and Keller, G. 2007. Identification and targeting of the ROSA26 locus in human embryonic stem cells. Nat. Biotechnol. 25:14771482. Kontgen, F. and Stewart, C.L. 1993. Simple screening procedure to detect gene targeting events in embryonic stem cells. Methods Enzymol. 225:878-890. Micallef, S.J., Janes, M.E., Knezevic, K., Davis, R.P., Elefanty, A.G., and Stanley, E.G. 2005. Retinoic acid induces Pdx1-positive endoderm in differentiating mouse embryonic stem cells. Diabetes 54:301-305. Micallef, S.J., Li, X., Janes, M.E., Jackson, S.A., Sutherland, R.M., Lew, A.M., Harrison, L.C., Elefanty, A.G., and Stanley, E.G. 2007. Endocrine cells develop within pancreatic bud-like structures derived from mouse ES cells differentiated in response to BMP4 and retinoic acid. Stem Cell Res. 1:25-36. Ng, E.S., Azzola, L., Sourris, K., Robb, L., Stanley, E.G., and Elefanty, A.G. 2005. The primitive streak gene Mixl1 is required for efficient haematopoiesis and BMP4-induced ventral mesoderm patterning in differentiating ES cells. Development 132:873-884. Nolden, L., Edenhofer, F., Haupt, S., Koch, P., Wunderlich, F.T., Siemen, H., and Brustle, O. 2006. Site-specific recombination in human embryonic stem cells induced by cell-permeant Cre recombinase. Nat. Methods 3:461-467. Pham, C.T., MacIvor, D.M., Hug, B.A., Heusel, J.W., and Ley, T.J. 1996. Long-range disrup-

tion of gene expression by a selectable marker cassette. Proc. Natl. Acad. Sci. U.S.A. 93:1309013095. Phelan, M.C. 2006. Techniques for mammalian cell tissue culture. Curr. Protoc. Mol. Biol. 74:A.3F.1-A.3F.18. Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: Somatic differentiation in vitro. Nat. Biotechnol. 18:399-404. Reubinoff, B.E., Pera, M.F., Vajta, G., and Trounson, A.O. 2001. Effective cryopreservation of human embryonic stem cells by the open pulled straw vitrification method. Hum. Reprod. 16:2187-2194. Sauer, B. 1993. Manipulation of transgenes by sitespecific recombination: use of Cre recombinase. Methods Enzymol. 225:890-900. Scacheri, P.C., Crabtree, J.S., Novotny, E.A., Garrett-Beal, L., Chen, A., Edgemon, K.A., Marx, S.J., Spiegel, A.M., Chandrasekharappa, S.C., and Collins, F.S. 2001. Bidirectional transcriptional activity of PGK-neomycin and unexpected embryonic lethality in heterozygote chimeric knockout mice. Genesis 30:259-263. Sidhu, K.S. and Tuch, B.E. 2006. Derivation of three clones from human embryonic stem cell lines by FACS sorting and their characterization. Stem Cells Dev. 15:61-69. Struhl, K. 2001. Subcloning of DNA fragments. Curr. Protoc. Mol. Biol. 13:3.16.1-3.16.2. te Riele, H., Maandag, E.R., and Berns, A. 1992. Highly efficient gene targeting in embryonic stem cells through homologous recombination with isogenic DNA constructs. Proc. Natl. Acad. Sci. U.S.A. 89:5128-5132. Thomason, L., Court, D.L., Bubunenko, M., Costantino, N., Wilson, H., Datta, S., and Oppenheim, A. 2007. Recombineering: Genetic engineering in bacteria using homologous recombination. Curr. Protoc. Mol. Biol. 78:1.16.11.16.24. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282:1145-1147. Urbach, A., Schuldiner, M., and Benvenisty, N. 2004. Modeling for Lesch-Nyhan disease by gene targeting in human embryonic stem cells. Stem Cells 22:635-641. van Deursen, J. and Wieringa, B. 1992. Targeting of the creatine kinase M gene in embryonic stem cells using isogenic and nonisogenic vectors. Nucleic Acids Res. 20:3815-3820. Vasquez, K.M., Marburger, K., Intody, Z., and Wilson, J.H. 2001. Manipulating the mammalian genome by homologous recombination. Proc. Natl. Acad. Sci. U.S.A. 98:8403-8410. Voytas, D. 2001. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Watanabe, K., Ueno, M., Kamiya, D., Nishiyama, A., Matsumura, M., Wataya, T., Takahashi, J.B.,

Generic Manipulation of Stem Cells

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Nishikawa, S., Nishikawa, S., Muguruma, K., and Sasai, Y. 2007. A ROCK inhibitor permits survival of dissociated human embryonic stem cells. Nat. Biotechnol. 25:681-686. Xu, R.H., Peck, R.M., Li, D.S., Feng, X., Ludwig, T., and Thomson, J.A. 2005. Basic FGF and suppression of BMP signaling sustain undifferentiated proliferation of human ES cells. Nat. Methods 2:185-190. Yasunaga, M., Tada, S., Torikai-Nishikawa, S., Nakano, Y., Okada, M., Jakt, L.M., Nishikawa, S., Chiba, T., Era, T., and Nishikawa, S. 2005. Induction and monitoring of definitive and visceral endoderm differentiation of mouse ES cells. Nat. Biotechnol. 23:1542-1550. Ying, Q.L., Stavridis, M., Griffiths, D., Li, M., and Smith, A. 2003. Conversion of embryonic stem cells into neuroectodermal precursors in adherent monoculture. Nat. Biotechnol. 21:183186. Zwaka, T.P. and Thomson, J.A. 2003. Homologous recombination in human embryonic stem cells. Nat. Biotechnol. 21:319-321.

Generation of hESC Reporter Knock-In Lines

5B.1.34 Supplement 11

Current Protocols in Stem Cell Biology

Guidelines for the Conduct of Human Embryonic Stem Cell Research

APPENDIX 1A

ABSTRACT This appendix provides a summary of and links to the ISSCR Guidelines for the Conduct of Human Embryonic Stem Cell Research and supporting documents. Curr. Protoc. Stem C 2009 by John Wiley & Sons, Inc. Cell Biol. 9:A.1A.1-A.1A.1.  Keywords: embryonic stem cells r guidelines r ISSCR research

The International Society for Stem Cell Research (ISSCR) calls for due consideration and appropriate oversight of human embryonic stem cell research to ensure transparent ethical and responsible performance of scientific experiments. These Guidelines, prepared by an ISSCR Task Force composed of international representatives, are meant to emphasize the responsibility of scientists to ensure that human embryonic stem cell research is carried out according to rigorous standards of research ethics, and to encourage uniform research practices that should be followed by all human stem cell scientists globally. The scope of these Guidelines includes principles for review and approval as well as oversight, enforcement mechanisms, procurement of materials with informed consent, principles for derivation, banking, and distribution of stem cell lines, dispute resolution, and ongoing review. There is an appendix with a glossary of scientific terms and a set of links to local, national, and international regulations related to embryonic stem cell research. These Guidelines are published at: http://www.isscr.org/guidelines/index.htm with links to Sample Consent documents for procuring materials, a sample Material Transfer Agreement (MTA), a Science magazine policy forum summarizing the guidelines, and biographies of the task force members.

Useful Information Current Protocols in Stem Cell Biology A.1A.1 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sca01as9 C 2009 John Wiley & Sons, Inc. Copyright 

A.1A.1 Supplement 9

ISSCR Guidelines for the Clinical Translation of Stem Cells

APPENDIX 1B

ABSTRACT This appendix provides a summary of and links to the ISSCR Guidelines for the Clinical Translation of Stem Cells and supporting documents. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 9:A.1B.1-A.1B.1.  Keywords: stem cells r guidelines r ISSCR clinical translation

In the fast-paced world of stem cell research, with the ever-present pressures for positive results that will translate to therapeutic use of human stem cells and their products, there is need for guidance and oversight to ensure that the scientific, clinical, regulatory, and social issues are properly addressed to guarantee that research results are appropriately translated into clinical therapies. These Guidelines define a roadmap for researchers and clinicians, outlining what needs to be accomplished to move stem cells from promising research to proven treatments for patients. They address issues related to cell processing and manufacture, preclinical studies, and clinical research, and contain 40 recommendations to ensure scientifically based, transparent, ethical, and medically and socially responsible clinical translation of stem cell–based therapies. Additionally, these Guidelines contain four appendices: the first is a Patient Handbook for those considering stem cell therapy; the second is a list of sites for local, national, and international regulations; the third is a list of related articles; and the fourth is a set of definitions. The Guidelines can be accessed at: http://www.isscr.org/clinical trans/index.cfm with links to the Patient Handbook (Appendix 1), additional resources (Appendix 2), a summary article from Cell Stem Cell, and the ISSCR and Cell Stem Cell press release.

Useful Information Current Protocols in Stem Cell Biology A.1B.1 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sca01bs9 C 2009 John Wiley & Sons, Inc. Copyright 

A.1B.1 Supplement 9

Standard Laboratory Equipment

APPENDIX 2

Listed below are pieces of equipment that are standard in the modern stem cell biology laboratory—i.e., items used extensively in this manual and thus not usually included in the individual materials lists. No attempt has been made to list all items required for each procedure in the Materials list of each protocol; rather, those lists note those items that might not be readily available in the laboratory or that require special preparation. See SUPPLIERS APPENDIX for contact information for commercial vendors of laboratory equipment. Applicator, cotton-tipped and wooden Autoclave Bag sealer Balances, analytical and preparative Beakers Bench protectors, plastic-backed (including “blue” pads) Biohazard disposal containers and bags Biosafety cabinet, tissue culture or laminar flow hood; filters air and maintains air flow pattern to protect cultured cells from investigator and vice versa Bottles, glass, plastic, and squirt Bunsen burners Centrifuges, low-speed (to 20,000 rpm) refrigerated, ultracentrifuge (20,000 to 80,000 rpm), large-capacity low-speed, tabletop, with appropriate rotors and adapters Centrifuge tubes and bottles, plastic and glass, various sizes Clamps Conical centrifuge tubes, plastic and glass Containers, assortment of glass and plastic, for gel and membrane washes Coplin jars, glass, for 25 × 75–mm slides Cryovials, sterile (e.g., Nunc) Cuvettes Desiccator and desiccant Dry ice Electrophoresis equipment, agarose and acrylamide, full-size and mini, with power supplies Film developing system and darkroom Filtration apparatus Forceps Fraction collector Freezers, −20◦ C, −70◦ C, and liquid nitrogen Fume hood Geiger counter Gel dryer Gloves, disposable plastic and heat resistant Graduated cylinders Heating blocks, thermostatically controlled for test tubes and microcentrifuge tubes

Hemacytometer and/or electronic cell counter Homogenizer Humidified CO2 incubator Ice bucket Ice maker Immersion oil for microscopy Lab coats Laboratory glassware Light box Liquid nitrogen Lyophilizer Magnetic stirrer, with and without heater, and stir bars Markers, including indelible markers, china-marking pens, and luminescent markers Microcentrifuge, Eppendorf-type with 12,000 to 14,000 rpm maximum speed Microcentrifuge tubes, 0.2-, 0.5-, 1.5-, 2-ml Microscope slides, glass, 25 × 75–mm, and coverslips Microscope with camera, upright, inverted, fluorescence, phase-contrast, dissecting Microtiter plate reader Mortar and pestle Ovens, drying and microwave Paper cutter, large Paper towels Parafilm Pasteur pipets and bulbs PCR thermal cycler and tubes pH meter pH paper Pipets, graduated Pipettors, adjustable delivery, 0.5- to 10-µl, 10- to 200-µl, and 200- to 1000-µl Polaroid camera or video documentation system Power supplies, 300-V for polyacrylamide gels, 2000- to 3000-V for other applications Racks, test tube and microcentrifuge tubes Radiation shield, Lucite or Plexiglas Laboratory Equipment

Current Protocols in Stem Cell Biology (2007) A.2.1-A.2.2 C 2007 by John Wiley & Sons, Inc. Copyright


A.2.1 Supplement 1

Radioactive waste containers for liquid and solid wastes Refrigerator, 4◦ C Ring stand and rings Rubber policemen or plastic scrapers Rubber stoppers Safety glasses Scalpels and blades Scintillation counter, β Scissors Shakers, orbital and platform, room temperature or 37◦ C Spectrophotometer, visible and UV range Speedvac evaporator Syringes and needles Tape, masking, electrician’s black, autoclave, and Time tape Test tubes, glass and plastic, various sizes, with and without caps

Timer Toolbox with common tools Trays, plastic and glass, various sizes Tubing, rubber and Tygon UV light sources, long- and short-wavelength UV transilluminator UV transparent plastic wrap (e.g., Saran Wrap) Vacuum desiccator Vacuum oven Vacuum supply Vortex mixers Waring blendor Water bath with adjustable temperature Water purification system X-ray film cassettes and intensifying screens

Standard Laboratory Equipment

A.2.2 Supplement 1

Current Protocols in Stem Cell Biology

SELECTED SUPPLIERS OF REAGENTS AND EQUIPMENT Listed below is contact information for commercial suppliers who have been recommended for particular items used in Current Protocols in Stem Cell Biology because: (1) the particular brand has actually been found to be of superior quality, or (2) the item is difficult to find in the marketplace. Consequently, this compilation may not include some important vendors of biological supplies. For comprehensive listings, see Linscott’s Directory of Immunological and Biological Reagents (Santa Rosa, CA), The Biotechnology Directory (Stockton Press, New York), the annual Buyers’ Guide supplement to the journal Bio/Technology, as well as various sites on the Internet.

Abcam 617-225-2272 Fax: 617-507-5831 http://www.abcam.com

Applied Biosystems 650-638-5800 Fax: 650-638-5998 http://www.appliedbiosystems.com

Bio-Rad Laboratories 800-424-6723 Fax: 510-741-1000 http://www.bio-rad.com

Curis 617-503-6500 Fax: 617-503-6501 http://www.curis.com

Addgene http://www.addgene.org

Applied Imaging 800-634-3622 Fax: 408-562-0250 http://www.aicorp.com

Calbiochem-Novabiochem 800-854-3417 Fax: 800-776-0999 858-450-9600 http://www.calbiochem.com

Dako Denmark A/S 45-44-85-95-00 http://www.dako.com

Affymetrix 888-DNA-CHIP (888-362-2447) Fax: 408-731-5441 http://www.affymetrix.com Agar Scientific 44-1279-813-519 Fax: 44-1279-815-106 http://www.agarscientific.com Agilent Technologies 877-424-4536 408-345-8886 Fax: 408-345-8474 http://www.home.agilent.com Amaxa See Lonza Ambion 512-651-0200 http://www.ambion.com American Pharmaceutical Partners 847-969-2700 888-391-6300 Fax: 800-743-7082 http://www.appdrugs.com American Type Culture Collection (ATCC) 800-638-6597 Fax: 703-365-2700 http://www.atcc.org Amersham Bioscience 732-457-8000 http://www.amersham.com Amgen 805-447-1000 Fax: 805-447-1010 http://www.amgen.com Amresco 800-829-2805 Fax: 440-349-1182 440-349-1199 http://www.amresco-inc.com/

Arctur See Molecular Devices Corporation ATCC See American Type Culture Collection Bando Chemical Industries http://www.bandousa.com Bard Parker See Becton Dickinson Baxter Healthcare 800-777-2298 Fax: 847-948-2000 http://www.baxter.com BD http://www.bd.com BD Biosciences 877-232-8995 http://bdbiosciences.com BD Falcon See BD Biosciences BD Pharmingen 800-848-6227 Fax: 858-812-8888 858-812-8800 http://www.pharmingen.com Beckman Coulter 800-742-2345 Fax: 800-643-4366 http://www.beckmancoulter.com Becton Dickinson Labware 888-237-2762 Fax: 800-847-2220 201-847-4222 http://www.bdfacs.com Bioline 888-257-5155 Fax: 775-828-0202 http://www.bioline.com Biomeda 800-341-8787 Fax: 650-341-8787 http://www.biomeda.com

Cambrex Corporation 201-804-3000 http://www.cambrex.com Canemco Inc. & Marivac Inc. 450-562-1451 Fax: 450-562-3430 http://www.canemco.com Cellomics 800-432-4091 412-770-2500 Fax: 412-770-2450 http://www.cellomics.com Chemicon International 800-437-7500 Fax: 909-676-8080 http://www.chemicon.com Clontech Laboratories 800-662-2566 Fax: 650-424-8222 http://www.clontech.com Cole-Parmer Instrument 800-323-4340 Fax: 847-247-2929 847-549-7600 http://www.coleparmer.com

Developmental Studies Hybridoma Bank 319-335-3826 http://dshb.biology.uiowa.edu/ DiaMed Lab Supplies Inc. 800-434-2633 Fax: 905-625-6280 http://www.diamed.ca Dow Corning 989-496-4400 Fax: 989-496-6731 http://www.dowcorning.com eBioscience 858-642-2058 Fax: 858-642-2046 http://www.ebioscience.com Eppendorf 800-645-3050 516-334-7500 Fax: 516-334-7506 http://www.eppendorf.com Ethicon See Johnson & Johnson Health Care Systems Falcon See Becton Dickinson Labware

Cooper Medical 64-9-3022955 http://www.coopermedical.co.nz

Fine Science Tools 800-521-2109 Fax: 650-349-1636 http://www.finescience.com

Corning and Corning Science Products 800-222-7740 Fax: 607-974-9000 http://www.corning.com

Fisher Scientific 800-766-7000 Fax: 412-562-8300 http://www3.fishersci.com

Costar See Corning Crown Scientific 61-1300-727-696 Fax: 61-1300-135-123 http://www.crownscientific.com.au

Fluka Chemical See Sigma-Aldrich GE Healthcare http://www.gehealthcare.com Gelaire http://www.gelaire.com.au

Suppliers

1 Current Protocols in Stem Cell Biology

CPSC Supplement 8

Gemini BioProducts 818-591-3530 http://www.gembio.com/ Gibco See Invitrogen Global Stem 888-545-0238, ext. 114 301-545-0238, ext. 114 Fax: 301-424-1989 http://www.globalstem.com Greiner Bio-One http://www.greinerbioone.com Hamilton Thorne Biosciences http://www.hamiltonthorne.com Hausser Scientific 215-675-7769 Fax: 215-672-9602 http://www.hausserscientific.com

Johnson & Johnson Health Care Systems 800-255-2500 Fax: 732-562-2212 http://www.ecatalog.ethicon.com Kendro Laboratory Products 800-522-SPIN Fax: 203-270-2166 203-270-2080 http://www.kendro.com Kent Scientific 888-572-8887 Fax: 860-567-4201 860-567-5496 http://www.kentscientific.com Kord-Valmark 800-452-9070 http://www.kord-valmark.com

Hunter Antisera 61-49620967 http://www.hum-molgen.org

Lafayette Instrument 800-428-7545 Fax: 765-423-4111 765-423-1505 http://www.lafayetteinstrument.com

HyClone Laboratories 800-HYCLONE Fax: 801-753-4584 http://www.hyclone.com

LEC Instruments 61-3-9763-0080 Fax: 61-3-9764-0086 http://lecinstruments.com

Illumina 800-809-4566 858-202-4566 Fax: 858-202-4766 http://www.illumina.com

Lonza http://www.lonza.com

Instrumedics 314-522-8671 Fax: 314-522-6360 http://www.instrumedics.com Integrated DNA Technologies 800-328-2661 319-626-8400 Fax: 319-626-8444 http://www.idtdna.com Invitrogen 800-955-6288 Fax: 760-603-7200 http://www.invitrogen.com Irvine Scientific 800-577-6097 Fax: 949-261-7800 http://www.irvinesci.com ISC BioExpress 800-999-2901 Fax: 801-547-5047 http://www.bioexpress.com The Jackson Laboratory 207-288-6000 http://www.jax.org John Morris Scientific http://www.johnmorris.com.au

Marienfeld 49-0-9343-6272-0 Fax: 49-0-9343-6272-25 http://www.superior.de/

MJ Research 800-PELTIER Fax: 617-923-8000 http://www.mjr.com

Qiagen 800-426-8157 Fax: 800-718-2056 http://www.qiagen.com

Molecular BioProducts 800-995-2787 Fax: 858-453-7551 http://www.mbpinc.com

Quality Biological 800-443-9331 301-840-8950 Fax: 301-840-0743 http://www.qualitybiological.com

Molecular Devices 408-548-6000 http://www.moldev.com Molecular Probes 800-438-2209 Fax: 541-465-8300 http://www.probes.com MP Biomedicals 800-854-0530 Fax: 800-334-6999 http://www.mpbio.com Nalge Nunc International 800-625-4327 Fax: 716-264-9346 http://www.nalgenunc.com Novocastra 44-0-191-215-0567 Fax: 44-0-191-215-1152 http://www.ebiotrade.com/buyf/ Novocastra/index.htm Orange Scientific 32-2-387-56-97 http://www.orangesci.com Pall http://www.pall.com

Matrix Technologies Corp. 800-345-2060 Fax: 603-595-0106 http://www.matrixtechcorp.com

Partec 49-2534-8008-0 Fax: 49-2535-8008-90 http://www.partec.com

MDS Nordion 800-465-3666 Fax: 613-592-6937 613-592-2790 http://www.mds.nordion.com

PeproTech 800-436-9910 Fax: 609-497-0253 http://www.peprotech.com

Mediatech/Cellgro 800-CELLGRO Fax: 703-471-0363 http://www.cellgro.com MidAtlantic Diagnostics 800-648-1151 Fax: 856-762-2000 http://www.midatlanticdiagnostics.com Millipore 800-645-5476 Fax: 800-645-5439 http://www.millipore.com Miltenyi Biotec 800-367-6227 Fax: 530-888-8871 http://www.miltenyibiotec.com

PerkinElmer 800-762-4000 http://www.perkinelmer.com

R & D Systems 800-343-7475 Fax: 612-379-2956 http://www.rndsystems.com Research Diagnostics 800-631-9384 Fax: 973-584-0210 973-584-7093 http://www.researchd.com Research Products International 800-323-9814 Fax: 847-635-1177 847-635-7330 http://www.rpicorp.com Roche Applied Science 800-262-1640 Fax: 800-428-2883 https://www.roche-applied-science.com Roche Diagnostics 800-262-1640 Fax: 317-845-2000 http://www.roche.com Sakura 800-725-8723 310-972-7800 Fax: 310-972-7888 http://www.sakuraus.com Santa Cruz Biotechnology 800-457-3801 Fax: 831-457-3800 http://www.scbt.com Henry Schein 631-843-5500 http://www.henryschein.com

Phoenix Biomedical Products 905-670-8299 Fax: 905-670-0195 http://www.phoenix-biomed.com

Serotec 44-1865-852722 Fax: 44-1865-373899 In the US: 800-265-7376 http://www.serotec.co.uk

Nicholas Piramal 91-22-3046-6666 http://www.nicholaspiramal.com

Sigma-Aldrich 800-358-5287 http://www.sigma-aldrich.com

Polysciences 800-523-2575 http://www.polysciences.com

Sorvall See Kendro Laboratory Products

Promega 800-356-9526 Fax: 608-274-4330 http://www.promega.com

Southern Biotechnology Associates 800-722-2255 Fax: 205-945-8768 205-945-1774 http://southernbiotech.com

Suppliers

2 CPSC Supplement 8

Current Protocols in Stem Cell Biology

Specialty Media 800-543-6029 Fax: 908-454-7774 http://www.specialtymedia.com

Taconic 888-TACONIC (888-822-6642) 518-697-3915 http://www.taconic.com

Thermo Electron 732-627-0220

Vectashield See Vector Laboratories

Thermo Scientific http://www.thermo.com

SpeedVac http://www.thermo.com

Techne 800-225-9243 Fax: 609-987-8177 609-452-9275 http://www.techneusa.com

Tree Star Software 800-366-6045 http://www.treestar.com

Vector Laboratories 800-227-6666 Fax: 650-697-3600 http://www.vectorlabs.com

Stem Cell Technologies 800-667-0322 Fax: 604-877-0713 http://www.stemcell.com Sterilin 44-0-844-844-3737 Fax: 44-0-844-844-2373 http://www.sterilin.co.uk Stoelting 920-894-2293 Fax: 920-894-7029 http://www.stoelting.com C. L. Sturkey 717-274-9441 Fax: 717-274-9442 http://www.sturkey.com Swemed See Vitrolife

Techno Plastic Products http://www.tpp.ch/ Ted Pella 800-237-3526 Fax: 530-243-2200 http://www.tedpella.com Tel-Test 281-482-2672 Fax: 281-482-1070 http://www.bioresearchonline.com/ storefronts/teltest.html Terumo Medical 800-283-7866 Fax: 732-302-3083 732-302-4900 http://www.terumomedical.com

Tyco Healthcare http://www.tyco.com United States Biological (US Biological) 800-520-3011 Fax: 781-639-1768 http://www.usbio.net Univentor 356-21-895824 Fax: 356-21-895835 www.univentor.com USA Scientific 800-LAB-TIPS Fax: 352-351-2057 352-237-6288 http://www.usascientific.com VacSax http://www.vacsax.plus.com

Vision BioSystems 800-753-7264 (North America) Fax: 781-610-1190 http://www.vision-bio.com Vitrolife 46 31 721 80 00 http://www.vitrolife.com VWR 800-932-5000 Fax: 610-431-9174 http://www.vwrsp.com Whatman 800-WHATMAN 973-245-8300 Fax: 973-245-8324 http://www.whatman.com Worthington Biochemical 800-445-9603 Fax: 732-462-3838 http://www.worthington-biochem.com

Suppliers

3 Current Protocols in Stem Cell Biology

CPSC Supplement 8