Current Protocols in Stem Cell Biology

Current Protocols in Stem Cell Biology Online ISBN: 9780470151808 DOI: 10.1002/9780470151808 Table of Contents 1. Forew...

Author: Mick Bhatia | Andrew Elefanty | Susan J. Fisher

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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.

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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

Preface

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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)

1A.1.2 Supplement 1


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


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.


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.


1A.1.5 Current Protocols in Stem Cell Biology

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


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



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


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


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.



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:


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


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


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


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


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



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.



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.



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


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.



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



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.



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


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



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



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


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



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.



Supplement 1

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


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.

1A.2.2 Supplement 1


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



Supplement 1

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.

1A.2.4 Supplement 1


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



Supplement 1

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.

1A.2.6 Supplement 1


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


Supplement 1

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.

1A.2.8 Supplement 1


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.



Supplement 1

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



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



Supplement 1

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



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.



Supplement 1

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



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



Supplement 1

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 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


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


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


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


Supplement 6

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.

1A.4.6 Supplement 6


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



Supplement 6

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.

1A.4.8 Supplement 6


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).



Supplement 6

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).



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.



Supplement 6

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.



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



Supplement 6

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.



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


Supplement 6

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.


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.



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


Supplement 6

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.


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.



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.



Supplement 6

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.


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.



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


Supplement 6

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, 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.



Supplement 6


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)


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.



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



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.


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.



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

1B.1.1 Supplement 2

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

1B.1.2 Supplement 2


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)

Characterization of Embryonic Stem Cells

1B.1.3 Current Protocols in Stem Cell Biology

Supplement 2

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 (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 (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


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).


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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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.



Supplement 9

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.


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.



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



Supplement 9

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.



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.



Supplement 9

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


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).



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.



Supplement 9

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


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.



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



Supplement 9

(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


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



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)



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)




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.



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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


continued



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.



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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)


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



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


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×


50 ml 2× SSC (see recipe) Add water to 500 ml and autoclave Store up to 6 months at −20◦ C



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.



Supplement 9


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)



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



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


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.



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.



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

1E.1.1 Supplement 1

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)

1E.1.2 Supplement 1


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 (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


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


1E.4.11 Current Protocols in Stem Cell Biology

Supplement 7

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


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)



Supplement 7

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).



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.



Supplement 7

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.



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.


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)



Supplement 7

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.

Isolation and Manipulation of Mouse Trophoblast Stem Cells


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.



Supplement 7

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.


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.



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.


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



Supplement 7

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


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.



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).



Supplement 7


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



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 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.

1F.4.4 Supplement 6


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


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.


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 .

1F.4.6 Supplement 6


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.


1F.4.7 Current Protocols in Stem Cell Biology

Supplement 6

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


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)

1F.4.8 Supplement 6


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


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


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)

1F.4.10 Supplement 6


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.



Supplement 6

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).


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).



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.



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


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.



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.



Supplement 6

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×


Combine the following: 80 g NaCl 2 g KCl 14.4 g Na2 HPO4 2.4 g KH2 PO4 800 ml H2 O continued



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.



Supplement 6

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.



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.



Supplement 6

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


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)


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


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


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).

1F.5.6 Supplement 6


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).


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.

1F.5.8 Supplement 6


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


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



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.



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


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.


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.



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 (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.

1F.6.6 Supplement 6


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 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)

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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.



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


PI + Ly6g/Mac1 PE single-stain control

A5


PI + B220 PE single-stain control

A6


PI + Ly1 PE single-stain control

B1


PI + GM cocktailb

B2


PI + B cell cocktailb

B3


PI + T cell cocktailb

B4


PI + GM cocktailb

B5



B6



C1

Mouse 1

PI + GM cocktailb

C2

Mouse 1


C3

Mouse 1


C4

Mouse 2

PI + GM cocktailb

C5

Mouse 2


C6c

Mouse 2


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.

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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.



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.


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



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., 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.



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.


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.



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



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


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



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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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

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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

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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


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

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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


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.

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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


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


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.

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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


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

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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.



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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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.



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


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



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


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.



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.



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


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.


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

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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.



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


Supplement 4

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 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.



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-gonadmesonephros (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.


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


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


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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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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-



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



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 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.

Placental Hematopoietic Stem Cells



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)



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.



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

Somatic Stem Cells



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.



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., Espié, 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.




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.


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


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.

Somatic Stem Cells


Supplement 3

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.


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).



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


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


Supplement 3

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.



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


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.

Somatic Stem Cells


Supplement 3

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.



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


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


Supplement 3

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


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


Supplement 3

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



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.

SUPPORT PROTOCOL 4

Somatic Stem Cells


Supplement 3

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



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.



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


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



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.


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 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

2C.1.12 Supplement 6


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).



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).


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×.



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.


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


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)



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 .



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.


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



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



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


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.



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.

Literature Cited Asahara, T., Murohara, T., Sullivan A., Silver, M., van der Zee, R., Li, T., Witzenbichler, B., Schattman G., and Isner J.M. 1997. Isolation of putative progenitor endothelial cells for angiogenesis. Science 275:964-967. Asahara, T., Masuda, H., Takahashi, T., Kalka, C., Pastore, C., Silver, M., Kearne, M., Magner, M., and Isner, J.M. 1999. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ. Res. 85:221228.


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.



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., Böhm, 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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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.

2C.2.2 Supplement 8


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).

2C.2.4 Supplement 8


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

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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

2C.2.6 Supplement 8



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


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 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

2D.1.4 Supplement 6


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


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


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.


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



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


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



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.

Somatic Stem Cells


Supplement 6

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

2D.2.4 Supplement 6

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


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)

Somatic Stem Cells


Supplement 6

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).

2D.2.6 Supplement 6


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

SUPPORT PROTOCOL 1 Somatic Stem Cells


Supplement 6

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 (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.


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.

Somatic Stem Cells


Supplement 6

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.



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.



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.




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.



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.



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).


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).



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.


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.



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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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



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.


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


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



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


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.



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.


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),



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.


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.



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.

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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 (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



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.

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Somatic Stem Cells


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Kanemura, Y., Mori, H., Kobayashi, S., Islam, O., Kodama, E., Yamamoto, A., Nakanishi, Y., Arita, N., Yamasaki, M., Okano, H., Hara, M., and Miyake, J. 2002. Evaluation of in vitro proliferative activity of human fetal neural stem/progenitor cells using indirect measurements of viable cells based on cellular metabolic activity. J. Neurosci. Res. 69:869879.

Kim, S.U. 2004. Human neural stem cells genetically modified for brain repair in neurological disorders. Neuropathology 24:159-171.

Kanemura, Y., Mori, H., Nakagawa, A., Islam, M.O., Kodama, E., Yamamoto, A., Shofuda, T., Kobayashi, S., Miyake, J., Yamazaki, T., Hirano, S., Yamasaki, M., and Okano, H. 2005. in vitro screening of exogenous factors for human neural stem/progenitor cell proliferation using measurement of total ATP content in viable cells. Cell Transplant. 14:673-682.

Kirschenbaum, B., Nedergaard, M., Preuss, A., Barami, K., Fraser, R.A., and Goldman, S.A. 1994. in vitro neuronal production and differentiation by precursor cells derived from the