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.



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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Wurmser, A.E., Palmer, T.D., and Gage, F.H. 2004. Neuroscience: Cellular interactions in the stem cell niche. Science 304:1253-1255. Yang, H., Mujtaba, T., Venkatraman, G., Wu, Y.Y., Rao, M.S., and Luskin, M.B. 2000. Regionspecific differentiation of neural tube-derived neuronal restricted progenitor cells after heterotopic transplantation. Proc. Natl. Acad. Sci. U.S.A. 97:13366-13371. Yasuhara, T., Matsukawa, N., Hara, K., Yu, G., Xu, L., Maki, M., Kim, S.U., and Borlongan, C.V. 2006. Transplantation of human neural stem cells exerts neuroprotection in a rat model of Parkinson’s disease. J. Neurosci. 26:1249712511. Yin, X.J., Ju, R., and Feng, Z.C. 2006. Experimental study on growth, proliferation and differentiation of neural stem cell from subventricular zone of human fetal brain at different gestational age. Zhonghua Er Ke Za Zhi 44:500504.

Zigova, T., Betarbet, R., Soteres, B.J., Brock, S., Bakay, R.A., and Luskin, M.B. 1996. A comparison of the patterns of migration and the destinations of homotopically transplanted neonatal subventricular zone cells and heterotopically transplanted telencephalic ventricular zone cells. Dev. Biol. 173:459-474. Zigova, T., Pencea, V., Betarbet, R., Wiegand, S.J., Alexander, C., Bakay, R.A., and Luskin, M.B. 1998. Neuronal progenitor cells of the neonatal subventricular zone differentiate and disperse following transplantation into the adult rat striatum. Cell Transplant. 7:137-156. Zimmerman, L., Parr, B., Lendahl, U., Cunningham, M., McKay, R., Gavin, B., Mann, J., Vassileva, G., and McMahon, A. 1994. Independent regulatory elements in the nestin gene direct transgene expression to neural stem cells or muscle precursors. Neuron 12:11-24.

Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S.,

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Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

UNIT 2D.4

Araceli Espinosa-Jeffrey,1 Dustin R. Wakeman,2, 3 Seung U. Kim,4 Evan Y. Snyder,3 and Jean de Vellis1 1

David Geffen School of Medicine at UCLA, Los Angeles, California University of California at San Diego, La Jolla, California 3 The Burnham Institute for Medical Research, La Jolla, California 4 University of British Columbia Hospital, Vancouver, British Columbia, Canada 2

ABSTRACT Here we document protocols for the production, isolation, and maintenance of the oligodendrocyte phenotype from rodent and human neural stem cells. Our unique method relies on a series of chemically defined media, specifically designed and carefully characterized for each developmental stage of oligodendrocytes as they advance from oligodendrocyte progenitors to mature, myelinating oligodendrocytes. Curr. Protoc. Stem Cell C 2009 by John Wiley & Sons, Inc. Biol. 10:2D.4.1-2D.4.26. Keywords: neural stem cells r NSC r oligodendrocyte specification r oligodendrocyte maturation r lineage progression r oligospheres r neurospheres

INTRODUCTION In this unit, protocols are provided for the derivation, expansion, and maintenance of the oligodendrocyte (OL) phenotype from both rodent and human neural stem cells (NSC). This unique method utilizes chemically defined media, each formulated and carefully characterized for specific developmentals stages of OL as they advance from OL progenitors (OLP) to mature myelinating OL (Fig. 2D.4.1; Neman and de Vellis, 2008). By providing hNSC with the nutrients specifically required at a particular moment in OL development, our system allows for the propagation of OL at a desired stage from OLP to mature premyelinating OL. Therefore, lineage progression can be manipulated by controlling the duration of a given developmental stage as needed, in a more “natural” manner, and without using gene transfer (Park et al., 2002b; Kim, 2004; Müller et al., 2006; Ahn et al., 2008), cocultures, or undefined substrates such as cell line–derived conditioned medium or animal serum.

Preparation of embryonic neural stem cells (NSC) The methodology described in this unit can be used to isolate and derive NSC lines from various species. Specific methods for the derivation of human NSC are detailed elsewhere (Svendsen et al., 1999; Villa et al., 2000; Palmer et al., 2001; Schwartz et al., 2003; Kim et al., 2006; De Filippis et al., 2007; Kim et al., 2008; Wakeman et al., 2009; also see UNITS 2D.2 & 2D.3). ISOLATION OF RODENT NEURAL STEM CELLS In this protocol, 1 to 14 embryos can produce a successful preparation because stem cells can be propagated many times to obtain the desired yield.

BASIC PROTOCOL 1

Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.4.1-2D.4.26 Published online September 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d04s10 C 2009 John Wiley & Sons, Inc. Copyright


stem cells embryonic

neural

NKX2.2 PSA-NCAM Nestin, VIM CD133, Pax6

OL progenitor

PDGFR Olig 1 Olig 2 NG2, GD3 A2B5, Sox9 NKX6, 1 NKX2.2

pre-OL

O4 Tf CNP GPDH PDGFR Olig 1 Olig 2

immature OL

mature OL

RIP GC O1 O4 Tf CNP GPDH

MBP, PLP GSTII, MOG RIP GC O1 O4 Tf CNP GPDH

GDM

OLDEM

SSeA-1 Oct4 Sox9 STM

OSM 20 hr

OSM 1,2 day

Figure 2D.4.1 Oligodendrocyte specification and lineage progression. Oligodendrocytes undergo sequential morphological changes as they develop from uncommitted NSC to a committed OLP and acquire characteristics inherent in a functional OL. The list of OL markers below each developmental stage is not exhaustive but represents frequently used markers to identify OLs and their developmental stage. Media (also see Reagents and Solutions): stem cell medium (STM; Espinosa-Jeffrey et al., 2002); OL specification medium (OSM; Espinosa-Jeffrey et al., 2002); glia defined medium (GDM; Espinosa de los Monteros and de Vellis, 1996); OLDEM (Espinosa de los Monteros et al, 1988, 1997). Modified from Arenander and de Vellis (1995).

Materials One timed-pregnant, embryonic day 14 to 16 (ED14 to ED16) Sprague-Dawley rat (Charles River Laboratories) Basal stem cell medium (STM-II; see recipe) supplemented with 1% (w/v) BSA (Sigma, cat no. A-3156) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) Complete stem cell medium (STMIIc; see recipe)


Dissection instruments, sterile (refer to numbers in Figure 2D.4.2): No. 1. Mayo scissors (Fine Science Tools, cat. no. 14010-17) No. 2. Lister scissors (Fine Science Tools, cat. no. 14131-14) No. 3. blunt-pointed forceps (Fisher, cat. no. 08-887) No. 4. iris scissors (Fine Science Tools, cat. no. 14060-09) No. 5. Moria iris forceps (Fine Science Tools, cat. no. 11373-12) No. 6. Dumont #7 forceps (Fine Science Tools, cat. no. 11297-10) No. 7. 140-μm and 230-μm sieves (Cellector, E-C Apparatus Corp.; http://www.thermo.com)



No. 8. 20-ml syringe (Kendall, cat. no. 520673; http://www.kendallhq.com) No. 9. 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 100 × 15–mm petri dish (bacterial grade, non TC-treated; BD Falcon, cat. no. 351029) 50-ml and 15-ml conical tubes Centrifuge (e.g., IEC Clinical) 100-mm anti-PSA-NCAM coated dishes (Support Protocol 1) 37◦ C, 4.5% CO2 incubator (adjustable to 5% if growth is slow), 95% humidity Additional reagents and equipment for isoflurane anesthesia of the mouse (UNIT 2A.5), assessing cell viability (Support Protocol 2), and counting cells using a hemacytometer (UNIT 1C.3) NOTE: All dissection instruments, plasticware, and glassware must be sterile. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals.

Collect the embryo brains 1. Prepare the work area and sterile tools in a biosafety hood (Fig. 2D.4.2). 2. Euthanize the rat by isoflurane inhalation (UNIT 2A.5). 3. Dissect and remove the uterus. Collect the placenta-containing embryos and place in basal stem cell medium containing 1% BSA at room temperature in a non-tissueculture-treated 100-mm-diameter petri dish. 4. Remove the embryos from their placenta. Place them in a 100-mm petri dish containing PBS at room temperature. Remove the cerebellum from each embryo.

7 8 filter 9 PBS 3

5

needle

Hank’s 6

4

2

1

Figure 2D.4.2 Instruments required for dissection. (1) Scissors for decapitation; (2) scissors to cut the head skin to expose the skull; (3) forceps to hold the head in place as you cut the skin and cut the skull cartilage with scissors (4) to expose the brain. Some users prefer the curved scissors (2) instead of (4). Use the same scissors to transfer the brain to the petri dish containing PBS. Forceps (5) and (6) are to hold the brain in place and remove the meninges, respectively. After removal of the meninges, place the brains in HBSS while dissecting the rest of the brains. A filter mesh (7) is used to filter the cell suspension after dissociation. A 20-ml sterile syringe (8) and sterile 18-G dissociation needle (9) are used to dissociate the cells.


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5. Dissect out the brain of each embryo and place in STM-II complete (STMIIc) medium at room temperature in a 100-mm non-tissue-culture-treated petri dish. 6. Separate the cortex from the rest of the brain and remove the meninges with forceps. Once devoid of meninges, combine cortex and pons (i.e., the rest of the brain).

Isolate the cells 7. Combine the tissue of all the brains without meninges in a 15-ml conical tube. Mechanically dissociate with the 18-G needle (attached to a 20-ml syringe) by gently aspirating the brain pieces (10 times) and releasing the suspension slowly with the needle against to the wall of the tube (try to minimize foaming). 8. Centrifuge 8 min at 450 × g, room temperature. Recover the supernatant with the cells in suspension and transfer to a 15-ml tube. 9. Add 2 to 4 ml of STMIIc to the chunks left over in the dissociation tube and dissociate again five to eight times. In place of steps 7 to 9, you can dissociate the cells for 2.5 min in a Stomacher 80 (Seward; http://www. brinkmann.com).

10. Filter the suspension of dissociated cells first through the 230-μm sieve and then through the 140-μm sieve to remove cell clusters. 11. Rinse the two sieves sequentially with 2 ml STM-II medium containing 1% BSA at room temperature, and add this medium to the tube containing the cells. 12. Collect the cells by centrifugation 8 min at 450 × g, room temperature. 13. Gently discard the supernatant. 14. Resuspend the cell pellet in 4 ml of complete stem cell medium (freshly prepared), and gently dissociate the pellet with the syringe and needle by aspirating it up and down twice.

Initiate the cultures 15. Assess cell viability (Support Protocol 1), count cells using hemacytometer (UNIT 1C.3), and plate onto fresh PSA/NCAM–coated dishes (2 × 106 cells/100-mm dish in 7 ml of STMIIc medium). 16. Incubate plated cells overnight at 37◦ C with 4.5% CO2 and 95.5% humidity. Younger cells do not yet express PSA-NCAM and will remain floating as small clusters, whereas the older cells will attach overnight.

Collect conditioned medium (CM) 17. On the next day, transfer the nonattached cells to a 15-ml conical tube, pellet cells by centrifugation for 8 min at 500 × g, room temperature, and remove and save the conditioned medium (CM). ◦

18. Collect, filter, and save the conditioned medium at 4 C for immediate use (or frozen for later use). CM is an excellent supplement to start NSC cultures from frozen stocks. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

19. Allow attached cells (see step 16) to grow to 70% to 90% confluency. 20. Resuspend the pellet from step 17 and dissociate in 4 ml of fresh STMIIc medium by passing through a 14-G needle eight times. Bring the volume to 8 ml with conditioned medium and plate on additional anti-PSA-NCAM coated plates. Alternatively, to dissociate the cells, place 1.0 ml of the cell suspension in a sterile 75-ml Erlenmeyer flask in 25 ml of STMIIc, and incubate with shaking at 37◦ C (Fig.2D.4.3).



E-16 cell suspension (mechanical dissociation) STMIIc

plate on anti-PSA-NCAM (coated dishes)

supernatant can be dissociated and re-panned

overnight

recover attached cells replate on fresh anti-PSA-NCAM

overnight

2-D cultures

NSC studies 3-D cultures

2-D

3-D attached on poly-D-lysine (well plates, chambers, dishes)

propagation

NSC

neurospheres

cryopreseration

neurospheres

Figure 2D.4.3 Rodent neural stem cell preparation. Following dissection, the cell suspension is plated on antiPSA-NCAM antibody–coated dishes and allowed to adhere. The process can be performed repeatedly to increase the numbers of neural stem cells, as two-dimensional cultures or three-dimensional “sphere” cultures (shown on the left side of the diagram). Every time cells are propagated, use anti-PSA-NCAM-coated dishes. Alternatively, cells can be propagated and immediately used for cell culture experiments (as shown on the right side of the diagram). While we prefer to use committed OL progenitors for cell transplants, other investigators also use uncommitted progenitors for grafting. Somatic Stem Cells


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21. Feed all cells every other day by removing 1/3 of the culture medium and adding the same volume of fresh STMIIc. 22. Switch the cells from 4.5% to 5.0% CO2 only if the cells are growing slowly. Leave them at 4.5% if the color of the medium stays red/orange. 23. Optional: To assess the phenotype of these cells, perform immunocytochemistry (Espinosa et al., 2002). SUPPORT PROTOCOL 1

PREPARATION OF ANTI-PSA-NCAM-COATED DISHES FOR SELECTING NSC BY IMMUNOPANNING The following method was developed based on published work (Wysocki and Sato, 1978; Williams and Gard, 1997) to isolate the rodent NSC population from the other cell populations in the brain during initial plating. We also use anti-PSA-NCAM coated dishes to propagate rodent NSC in two-dimensional cultures (Espinosa-Jeffrey et al., 2002). Please refer to the literature for specific methods on the selection of human NSC during primary derivation (Wakeman et al., 2009). We have chosen immunopanning as opposed to flow cytometric cell sorting because we find that cell survival approaches 100% when selecting the desired cell type via immunopanning. We know from both the experience of other scientists and our own that the survival rates are never this high when using FACS. Moreover, immunopanning can be performed in the standard culture vessel and is as simple as plating the cells on the adequate substrate for cell selection.

Materials 50 mM Tris·Cl, pH 9.5 Bovine serum albumin (BSA; Sigma, cat. no. A-3156) Anti-PSA-NCAM antibody (Iowa DSHB, http://dshb.biology.uiowa.edu/, cat. no. 5A5) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) 100 × 15–mm petri dish (bacterial grade, non-TC-treated; BD Falcon, cat. no. 351029) 1. Prepare the immunopanning cocktail:

50 mM Tris·Cl, pH 9.5 containing: 1% (w/v) BSA 50 μg/ml anti-PSA-NCAM antibody. 2. Coat the bottom surface of 100-mm non-tissue-culture-grade petri dishes by adding 4 to 5 ml per dish of the anti-immunopanning cocktail and incubating 30 min at 37◦ C. Flasks may also be coated by this protocol: use 2.5 ml to coat a 12.5-cm2 flask or 5 ml to coat a 75-cm2 flask.

3. Remove the cocktail. Wash petri dishes three times, each time with 5 ml PBS, then once with 5 ml PBS containing 1% BSA just before using. Do not allow the plates to dry. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

4. Cover extra plates (still containing the PBS/BSA) with foil and store at 4◦ C for up to 10 days.



ASSESSING CELL VIABILITY Cell viability can be determined with the SYTOX blue nucleic acid stain (Molecular Probes/Invitrogen). Cells with compromised plasma membranes are labeled by SYTOX binding to nucleic acids and detected by fluorescence microscopy.

SUPPORT PROTOCOL 2

Materials Tris-buffered saline (TBS; see recipe) Phosphate-buffered saline (PBS; Sigma, cat. no. P-5368) 1 μM SYTOX blue nucleic acid stain (Invitrogen Molecular Probes, cat. no. S7020) in PBS Microscope slides and coverslips Fluorescence microscope 1. Gently wash cells three times, each time with 5 ml TBS. 2. Harvest cells with a cell scraper and transfer them to a 15-ml tube. Resuspend in 2 ml PBS. Do not use enzymes.

3. Add 1 μl of 1 μM SYTOX to each tube (final concentration, 5 nM). 4. Incubate 12 min. 5. Remove the solution and wash the cells five times, each time with 5 ml TBS, centrifuging 3 min at 500 × g, room temperature, each time. 6. Resuspend the cells in 1 ml/tube of PBS. Take an aliquot and add a drop to a microscope slide. Add a coverslip and examine with a fluorescence microscope. 7. Determine the number of positive cells per random field in a fluorescence microscope and record as a percentage of the total number of cells in the field.

PROPAGATION OF RODENT NSCs AS TWO-DIMENSIONAL CULTURES NSCs can be propagated in two-dimensional (2-D) or three-dimensional (3-D) cultures. When attached, NSCs (2-D cultures) tend to grow faster and are therefore ideal for creating a large cell stock quickly before starting specific studies. In addition, we have developed a new method for expansion and maintenance of human NSC in NB-B-27 medium (see Reagents and Solutions) as multilayer adherent network (MAN) cultures, with increased proliferation rates compared to standard sphere-forming assays (UNIT 2D.3). In order to accommodate the difference in basal media, human NSC can either be initially derived in STMIIc media (replacing NB-B-27), or previously established hNSC cultures may be slowly transitioned away from the basal NB-B-27. Simply substitute 25% STMIIc for 1 week, followed by successive weeks at 50%, 75%, and finally 100% STMIIc after 1 month.

BASIC PROTOCOL 2

This procedure can be used every time cells are replated.

Materials Cultures of freshly isolated neural stem cells (Basic Protocol 1) or their progenitors Hanks’ buffered salt solution (HBSS) without Ca2+ or Mg2+ Complete stem cell medium (STMIIc; see recipe) Cell Freezing Medium, serum-free, 1× (Sigma, cat. no. C2639) Cell scraper 15-ml conical tubes

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Centrifuge (e.g., IEC Clinical) 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 12.5-cm2 and 75-cm2 tissue culture flasks (Falcon), anti-PSA-NCAM-coated (Support Protocol 1) 1.2-ml cryovials Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1C.3) and freezing cells (Support Protocol 3) NOTE: All steps are performed at room temperature (20◦ C).

Collect the cells 1. When confluency of the neural stem cells has been reached, remove the supernatant conditioned medium (CM; save for step 4), add 5 ml of HBSS without Ca2+ or Mg2+ , detach the cells with a cell scraper, and transfer to a 15-ml conical tube (accommodating cells from one to three dishes). 2. Rinse the dish once with 2 ml HBSS (without Ca2+ or Mg2+ ), add it to the tube, and centrifuge 8 min at 450 × g. Discard the supernatant. 3. Resuspend the cell pellet in 3 ml STMIIc, dissociate gently using 18-G needle and syringe, and centrifuge 5 min at 450 × g. 4. Resuspend cells in 2 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM (use CM from step 1). Replate cells on anti-PSA-NCAM-coated plates as described in Basic Protocol 1. If you have repeated this process several times and the cell pellet is 0.5 ml volume or larger, divide the cell suspension into two parts. One part of the suspension will be used to start a frozen stock (see Support Protocol 3). The second half of the cell suspension is further dissociated using a needle (as described in Basic Protocol 1; however, the sieves (used in step 10 of Basic Protocol 1) are not necessary (single-cell suspension is obtained using needle and syringe), and replated as described below.

Replate the cells 5. Count the number of cells/ml and adjust the volume to 15 ml with a freshly prepared mixture of 2 parts STMIIc and 1 part CM (use CM from step 1). 6. Plate 2 ml of the cell suspension in each of five 12.5-cm2 cell culture flasks coated with anti-PSA-NCAM.

Passage the cells 7. Feed cells with a freshly prepared mixture of 2 parts STMIIc and 1 part CM every other day until they reach 80% to 90% confluency, and repeat steps 1 to 7 to increase the number of cells. 8. When four or more 12.5-cm2 flasks reach confluency, harvest the cells as described above, and seed the equivalent content of cells from three 12.5-cm2 flasks into one 75-cm2 flask coated with anti-PSA-NCAM. Feed the cells with a freshly prepared mixture of 2 parts STMIIc and 1 part CM in a total volume of 10 ml/flask. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

9. After 1 to 2 days, check to see if the culture medium is red. If the medium is turning orange, add 3 ml of STMIIc. Repeat this step as needed. Add 3 ml of STMIIc every day only if the medium changes color. If cells seem not to grow, but look healthy, or if the culture medium is not red but purple, you will need to remove 1/2 of the plating medium and bring the volume up to 10 ml with a freshly prepared mixture of 2 parts STMIIc and 1 part CM.



If the opposite is true and the culture medium turns orange overnight, the cells have proliferated heavily, and you will need to replace the culture medium and seed more 75-cm2 flasks (one 75-cm2 flask per three 12.5-cm2 flasks).

10. Optional: When cells reach confluency, freeze the contents of one 75-cm2 flask (Support Protocol 3) in 1 ml of freezing medium in a cryovial. When propagating cells to create frozen stocks, we strongly recommend maintaining a “mother flask” by scraping most, but not all of the cells attached to the flask. After removing the detached cells, feed the mother flask with a 1:1 mixture of fresh STMIIc and CM to ensure continuity of these cultures (in case replated cells do not look healthy, grow slowly, or die).

11. Optional: After accumulating at least 10 to 15 vials of cryopreserved NSC in a frozen stock, grow NSCs as neurospheres (three-dimensional; see Alternate Protocol) for slower growth, allowing more time to devote to experiments. While cells are floating, even in STMIIc their metabolism seems slower, but if replated they behave normally.

FORMATION, PROPAGATION, AND MAINTENANCE OF NEUROSPHERES IN THREE-DIMENSIONAL CULTURES

ALTERNATE PROTOCOL

Suspension aggregate, or “neurosphere,” three-dimensional cultures are an alternative strategy to propagate NSCs at a slower pace than that of attached cells, while preserving most of the standard characteristics of a proper NSC. NSC suspension cultures are started from freshly dissociated NSC two-dimensional cultures (Basic Protocol 2) and grown in sterile Erlenmeyer flasks to prevent attachment and encourage free-floating spherical growth.

Materials Conditioned medium (see Basic Protocol 2) Complete stem cell medium (STMIIc; see recipe) Established NSC cultures (Basic Protocol 2) Glass Erlenmeyer flasks, 25-ml or 50-ml with cap 37◦ C incubator with shaker 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 50-ml conical tubes Centrifuge (e.g., IEC Clinical) 0.22-μm sterile filters NOTE: All steps are performed at room temperature (20◦ C). 1. After harvesting cells as described in Basic Protocol 2, resuspend the cells from one 75-cm2 flask in 6 ml fresh STMIIc. 2. Place 15 ml of fresh STMIIc and 3 ml of conditioned medium (CM) into a 25- or 50-ml Erlenmeyer flask (depending on number of cells). 3. Add 2 ml of the cell suspension (freshly harvested from two-dimensional cultures as described in Basic Protocol 2), and close the top of the flask partially to allow for O2 /CO2 exchange. Thus, one 75-cm2 flask will result in three Erlenmeyer flasks of neurospheres. Somatic Stem Cells


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4. Place the flask, continuously shaking at 90 rpm, in the incubator. If placing a shaker in the incubator is not an option due to safety regulations, the cell suspension may be placed directly into two noncoated petri dishes (bacterial grade) to prevent cell attachment.

5. Add 1.5 ml fresh STMIIc, every other day, and dissociate routinely (three times gently) with a syringe and needle in the same flask (as described for pellet dissociation in Basic Protocol 2) to keep the spheres at a small size. This process allows for increased sphere formation without the negative potential for spontaneous differentiation. It also allows cells more exposure to the fresh nutrients in the culture medium, helping preserve “stemness” in all cells.

6. When the culture medium turns orange overnight, collect the contents of the Erlenmeyer flask with a pipet and place into one 50-ml conical tube. Centrifuge for 6 min at 450 × g. 7. Slowly collect the CM (supernatant), filter (0.22-μm), and save for replating cells. 8. Resuspend the pellet in 4 ml of STMIIc to dissociate larger spheres. Add 21 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM. At this point, the neurospheres should all be easily dissociated. If some spheres remain large in spite of repeated dissociation, use the sieves (see Basic Protocol 1 materials list) to eliminate the large clumps, instead of vigorously dissociating them. This step will prevent significant cell death at the time of replating.

9. Seed cells on desired containers for experiments, or continue to propagate NSCs as two- or three-dimensional cultures (see Basic Protocol 2). SUPPORT PROTOCOL 3

CRYOPRESERVATION/THAWING OF NSC STOCKS We recommend collecting cells for frozen stocks at low passage number. Human NSC are cryopreserved using modified methods found elsewhere (Wakeman et al., 2009). In addition, the method formerly described for rat and mouse NSC (Espinosa-Jeffrey et al., 2002) can also be used to stock human NSC. The present methods are to be used for basic research and can be optimized for translational research if such cells are approved for the clinic. Therefore, we recommend the use of serum-free freezing medium as well as all other animal-free components as indicated within this unit. It would defeat the purpose to perform the full preparation and maintenance utilizing animal-free products and then expose the cells to fetal bovine serum or other types of sera at the moment of cryopreservation.

Materials NSC cultures ready for freezing (Basic Protocol 2) Hanks’ balanced salt solution (HBSS) without Ca2+ or Mg2+ Complete stem cell medium (STMIIc; see recipe) Cell Freezing Medium, serum-free, 1× (Sigma, cat. no. C2639) Liquid nitrogen Conditioned medium (CM; see Basic Protocol 2) Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

Cell scrapers Centrifuge (e.g., IEC Clinical) 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) 1.2-ml cryovials



Cryogenic slow-freezing chamber (Nalgene, cat. no. EW-44400-00) 2-ml tubes (Fisher) Anti-PSA-NCAM coated tissue culture vessels (Support Protocol 1) Additional reagents and equipment for testing cell viability (Support Protocol 2) NOTE: All steps are performed at room temperature (20◦ C).

Collect and freeze the NSC 1. Allow NSCs to grow to 70% to 90% confluency. Remove all of the cell culture medium, add 5 ml of HBSS (without Ca2+ or Mg2+ ) to each 100-mm petri dish or 10 ml to each 75-cm2 flask, and detach cells by gently scraping the culturing surface. 2. Centrifuge the cells 8 min at 450 × g, and resuspend in 3 ml STMIIc medium. 3. Gently dissociate cells using an 18-G needle and syringe, centrifuge 8 min at 450 × g, and discard the supernatant. 4. Gently resuspend the pellet from one 100-mm petri dish or 75-cm2 flask in 1 ml of serum-free freezing medium. 5. Transfer the contents to a 1.2-ml cryovial, and place the vial(s) in a cryogenic freezer container overnight for slow freezing. 6. Next day, place the vials in liquid nitrogen for long-term storage.

Thaw NSCs 7. To reanimate NSCs, defrost cryovials quickly in a 37◦ C water bath, and transfer the contents of the vial to a 2-ml tube containing 1 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM at 37◦ C. 8. Centrifuge gently 5 to 7 min at 350 × g. 9. Remove the supernatant, add 1 ml of a freshly prepared mixture of 2 parts STMIIc and 1 part CM at 37◦ C, resuspend the cell pellet, and remove a small aliquot to test the initial cell viability. 10. Count the number of viable cells in the tube (∼1 × 106 cells expected) as described in Support Protocol 2. 11. Plate cells onto an anti-PSA-NCAM-coated vessels (petri dishes or 75-cm2 flasks, plate the equivalent of 1 vial/75 cm2 flask). If the yield is lower, use 25-cm2 flasks to increase the cell density necessary for healthy growth. Seeding low-density cultures in large containers decreases the proliferation rate and might be detrimental to the culture.

12. To propagate NSCs after replating, proceed as described in Basic Protocol 2. Alternatively, when cells have reached 90% confluency, either freeze them or use them for experiments.

OLIGODENDROCYTE COMMITMENT IN TWO- AND THREE-DIMENSIONAL CULTURES

BASIC PROTOCOL 3

During development, the nutritional and environmental needs of cells change as they lose multipotency and become lineage restricted. The present system is based on the modification of nutrients contained in the cell culture medium and the percentage of CO2 needed to optimize and direct lineage restriction towards the oligodendrocyte (OL) phenotype. Like NSCs, OL progenitors (OLPs) can be propagated in two- and threedimensional cultures. When attached (two-dimensional cultures), OLPs grow faster and,

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thus, two-dimensional cultures are ideal to create an OLP cell stock quickly before starting specific in vitro cell culture or in vivo transplantation studies. A diagram of the following steps can be found in Figure 2D.4.4.

Materials NSC cultures (2-D or 3-D; Basic Protocol 2 or Alternate Protocol) Hanks’ balanced salt solution (HBSS) without Ca2+ or Mg2+ OL specification medium (OSM-II; see recipe) Cell scraper 15-ml conical tubes 20-ml syringe 18-G Quincke spinal luer-lock needle for dissociation (100-mm length; Unimed; http://www.unimed.ch/) Anti-IgM coated 100-mm petri dishes or tissue culture flasks: prepare as in Support Protocol 1 but substitute goat anti-IgM antibody (ABR, sold by Thermo Scientific, cat. no. PA1-86106) for anti-PSA-NCAM antibody 37◦ C, 4.5% CO2 incubator 12-ml syringe (Tyco Healthcare, cat. no. 512852) Additional reagents and equipment for maintaining cells (Basic Protocol 1) NOTE: All steps are performed at room temperature (20◦ C). NOTE: We recommended precalibrating the percentage of CO2 1 day before plating the cells. If the incubator is shared with other people or needed at 5% for NSC propagation and maintenance, we recommend using tissue culture flasks for 2-D cultures instead of petri dishes. Close the cap of the flask completely and then open it one-quarter of a turn before placing in the incubator at 5% CO2 . For propagation and maintenance of OL spheres, the Erlenmeyer flask should also be kept open just enough to ensure O2 /CO2 exchange. When using 4.5% CO2 , loosen the caps of the flasks until half-way open. 1. When NSCs reach confluency, remove the supernatant (CM; save for use in subsequent steps), and add 5 ml of HBSS without Ca2+ or Mg2+ . 2. Detach the cells with a cell scraper, transfer into a 15-ml tube (which accommodates one to three petri dishes), rinse once with 2 ml of HBSS (without Ca2+ or Mg2+ ), and centrifuge 8 min at 450 × g. 3. Resuspend the cell pellet in 3 ml OSM-II medium and gently dissociate three times using a syringe and 18-G needle. Centrifuge 8 min at 450 × g to pellet the cells.

For two-dimensional OL cultures 4a. Resuspend the cells in a freshly prepared 1:1 mixture of OSM-II and CM. Seed cells on anti-IgM coated dishes or flasks. By using anti-IgM unconjugated antibody for coating, not only PSA-NCAM-positive cells will attach to the plate or dish, but also the cells that begin to express A2B5+ gangliosides. The panning strategy can be used at later stages to select OL cells at a single developmental stage (see Fig. 2D.4.1), e.g., pre-OL, which can be selected using anti-O4. Culture System for Rodent and Human Oligodendrocyte Specification, Lineage Progression, and Maturation

The CM used here and in the following step is self-conditioned STMIIc.

5a. Maintain the cells as described in Basic Protocol 1 but using a mixture of 2 parts OSM-II and 1 part CM. From this point on, maintain the CO2 concentration in the incubator at 4.5%. Cells switch from NSCs to OLP (third stage shown in Fig. 2D.4.1) within 20 hr in contact with OSM, at which time the cells start to express transferrin (Tf).



dissociation NSC in STM/OSM 2:1 2-D cultures

3-D cultures CM/OSM 2:1

overnight

slow propagation (oligospheres)

CM/OSM 1:1

fast propagation: plate on anti-PSA-NCAM

overnight

100% OSM

slow propagation

fast propagation

in vitro studies cryopreservation GDM

transplant studies

Figure 2D.4.4 Oligodendrocyte specification. The transition of NSC to commit to the OL lineage is brief, but sequential rather than abrupt. In order for cells to survive, they must acclimate to their new environment. OLP can be propagated to create frozen stocks as three-dimensional “oligosphere” cultures (shown on the left of the diagram) or frozen without propagation (as shown in the sequence in the center of the diagram). OLP can also be propagated in two-dimensional cultures for cryopreservation, for specific cell culture experiments, or for cell replacement therapies (as shown on the right side of the diagram).

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6a. Feed the cells with a mixture of 2 parts OSM-II and 1 part CM (i.e., self-conditioned OSM-II) every other day until they reach 80% to 90% confluency (3 to 5 days). This process can be repeated several times to attain a large number of cells for freezing (if desired).

For three-dimensional OL cultures 4b. Alternatively, to grow OL spheres to create/enrich a frozen stock of OLP, place the equivalent of 2 mm2 of cells (pellet size after cells are dissociated and in suspension) in a 25-ml Erlenmeyer flask with 15 ml of a mixture of 2 parts (10 ml) OSM-II and 1 part (5 ml) CM (i.e., self-conditioned OSM-II). If the pellet is 4 mm2 (∼12-15 × 106 OLP), use a 50-ml Erlenmeyer flask. Prepare the cell suspension and place in a total volume of 25 ml of a mixture of 2 parts OSM-II and 1 part CM (i.e., self-conditioned OSM).

5b. Feed OL spheres with fresh OSM-II every other day by adding 3 ml of freshly prepared OSM-II (no CM). 6b. When spheres start to become larger than 2 mm, gently dissociate by aspirating them one to two times in the same flask with the 18-G needle using a 12-ml syringe (sterile). 7. When the culture medium starts to turn orange, recover and centrifuge the spheres from one flask, and split cells into more Erlenmeyer flasks (1 to 4). These may be used for experiments or cryopreserved as previously described (Support Protocol 3). BASIC PROTOCOL 4

CULTURING OLIGODENDROCYTES FOR LINEAGE PROGRESSION AND MATURATION The nutritional needs for a committed cell within the OL lineage differ considerably as the cells progress and mature to the next developmental stage. These cells need to start synthesizing enzymes and proteins related to myelination; therefore, the energy demand is enormous compared to their earlier stage where migration and proliferation are the basic functions. The culture medium “GDM” (glial defined medium) was first designed to maintain 04+ , GC+/− , CNP+/− cells (for details. see “pre-OL” in Fig. 2D.4.1). Later, we realized that GDM also induced the transition of OLP to pre-OL (Espinosa de los Monteros et al., 1997). OL can be sustained at a given developmental stage by keeping them in one of the stage-specific culture media described here. Not all cell types offer the possibility for studying commitment and full differentiation when grown in culture, and one of the best examples comes from glial biology. Astrocytes grown in artificial cell culture conditions have allowed us to understand many of their functions and interactions with neurons and oligodendrocytes and how they play an integral part in mediating disease pathology; however, to our knowledge, there is no definitive proof of a fully matured, terminally differentiated astrocyte that can be studied throughout its progression and maturation in cell culture. In contrast to astroglial biology, we appear now to have the necessary tools to terminally differentiate oligodendrocytes in vitro, which even produce large amounts of compact myelin-like membranes in the absence of axons.


Most cell culture methods for maintaining OL only allow the researcher to address commitment, survival, or maintenance in a cell type that will, by default, tend to progress from the progenitor stage into a more mature stage uncontrollably (i.e., beyond control by the researcher) when maintained in a fairly rich environment conducive to and promoting myelinogenic properties. However, in our model, the design of several culture media specific for multiple OL developmental stages provides us with the ability to control



lineage progression of normal OL at multiple lineage transitions. In addition, these subtype-specific media allow us to determine potential deficiencies in diseased or stressed OL derived from transgenic/mutant animals and tissues donated by human subjects. Therefore, development of OL-lineage-specific media formulations allows us to further model disease mechanisms and determine how they affect OL at different stages of development. Furthermore, we can then use these data to design specialized culture media aimed at either further protecting the cell or preventing specific mechanisms from potentially occurring in OL-related diseases, or as part of their inherent injury response. Utilizing this platform, we can apply high-throughput small-molecule libraries to our defined media and determine how specialized formulations may effectively aid in the restoration and repair of degenerating tissue.

Materials OLPs (Basic Protocol 3, step 6a) OL specification medium (OSM-II; see recipe) GDM medium (see recipe) Recombinant human basic fibroblast growth factor (bFGF; Invitrogen) OLDEM medium (see recipe) Poly-D-lysine-coated wells/plates (see recipe) Additional reagents and equipment for oligodendrocyte differentiation in two-dimensional culture (Basic Protocol 3) Culture for pre-OLs 1. In order to obtain pre-OL (along the OL lineage), plate OLPs using OSM-II (as in Basic Protocol 3). As in previous steps, they may be propagated as OL spheres (three-dimensional) or as twodimensional cultures on anti-IgM coated flasks or petri dishes, or directly on cell-culture grade plastic. See Figure 2D.4.4 for options.

2. On the next day, remove one-half of the volume of the plating medium (OSM-II) and add the same volume of GDM. Continue incubation for a minimum of 2 days, or until 90% confluence is reached. 3. To obtain more OLP/pre-OL, grow cells as two- or three-dimensional cultures in the presence of bFGF (Fig. 2D.4.5). To keep progeny cells at the same stage as the parent cells, add 2 ml fresh GDM containing 20 ng/ml bFGF every other day until 90% confluence is reached. For cell replacement therapies, we suggest using cells at this stage (1 to 2 days after plating without bFGF; see alternatives described in Fig. 2D.4.4), as cells are still highly motile and readily migrate within the host post-natal and/or adult rodent brain and/or spinal cord.

4. To enhance maturation of cells into the next developmental stage, culture OL as two-dimensional cultures (Basic Protocol 3). Plate 1 × 105 cells/ml in GDM for at least 2 days (if plated in GDM without bFGF), or 4 days (if plated in GDM with bFGF) without further bFGF supplementation (Fig. 2D.4.5). After exposure to GDM, cells express myelin enzymes and proteins, and they display multipolar, branched cell processes, but not a myelin-like membrane. In addition, OL maintained in GDM for at least 4 days (without bFGF or any other factors) can be further induced to a fully mature myelinating stage. Also see Figure 2D.4.6.

5. To obtain fully mature OL, plate as two-dimensional cultures (Basic Protocol 3) at 1 × 105 cells/ml onto poly-D-lysine coated wells/plates or uncoated petri dishes in 10 ml of a 1:1 mixture of GDM and OLDEM (OL maturation medium). Culture for 1 to 5 days, then replace with 100% OLDEM for further culture (Fig. 2D.4.5).

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GDM bFGF 4 days

bFGF overnight

OLP propagation

maturation

bFGF overnight

bFGF overnight oligospheres OLDEM 1:1 1 to 5 days

GDM bFGF 2 days

bFGF overnight cocultures myelination studies

OLDEM 100% transplant studies

cryopreservation

mature myelinating

Figure 2D.4.5 Oligodendrocyte lineage progression and maturation. After commitment of NSC to the OL lineage, cells are propagated at the OLP stage to create a frozen stock (steps indicated on the left portion of the flow chart) or processed further for transplantation studies (as shown by the middle arrow on the diagram). To allow OLP to further mature along the OL lineage and become myelinated, cells are transitioned into OLDEM for at least 48 hr. Once OL have reached this stage of maturation, they are excellent for cell culture studies but are not recommended for cell grafting. Detachment from the substrate can damage the numerous delicate cell processes; therefore these cultures are no longer a quality source for cell transplants.

6. Every 4 days, feed the cells by replacing all of the culture medium with fresh OLDEM. Myelinating OL express myelin markers (see Fig. 2D.4.1, last two columns). The medium should look red, not orange. If it turns orange, add more medium while feeding the cells.


These cells will express myelin enzyme levels comparable to those found in pure myelin within 5 days after having been introduced to 100% OLDEM. As they mature, cells will synthesize myelin-like membranes in vitro even in the absence of neurons. They can be maintained for a number of weeks if they are kept subconfluent; however, if the culture becomes overcrowded, cells will deteriorate and die.



B

C

OSM 2 days

OSM 3 days

GDM 1 day

DD

EE

F

Nestin Tf Tf

N estin T Nestin Tff

O 4 /MBP O4

G

H H

I

HuOLP, OSM 2 days

GDM 10 days, MBP

HuOL/Rat Neurons

A

Figure 2D.4.6 Phase-contrast view of neural stem cells derived from embryonic day 16 rat brain at passage number 2 (P2). NSC were plated and maintained in OSM for 2 days (A), 3 days (B), or 3 days in OSM then switched to GDM for 1 day (C). Cells in OSM still proliferate while in OSM. When cells from (A or B) are plated and maintained in OSM on poly-D-lysine-coated coverslips for 1 day, they start to display a bipolar or multipolar morphology (D) and most express the immature precursor marker, nestin (green) but not Tf (red), an early marker for OL. After 2 days in OSM, bipolar nestin+ cells coexpress transferrin (Tf) (E). After 4 days in OSM, cells were switched to GDM for 1 day; they developed numerous cell processes and coexpressed sulfatides (recognized by the anti-O4 antibody, green) and myelin basic protein (MBP; red) (F). Panels G to I are human cells. (G) Phase-contrast view of human NSCs (HFB-2050) acclimated and expanded in STM, then replated and maintained in OSM for 2 days. (H) OL derived from human NSCs (HFB-2050) were specified to the OL lineage with OSM and maintained in GDM for 10 days. OL matured and started to express MBP (red). (I) Rat cortical neurons (NFM-200-red) were cultured for 10 days, then human OLP derived from NSC (HFB-2050) were added in coculture for 24 hr. These cells were labeled with human nuclei marker (HuNu, green).

PROPAGATION OF OLIGODENDROCYTES FOR IN VITRO MYELINATION ASSAYS

BASIC PROTOCOL 5

To perform myelination studies in vitro, it is recommended to start with OL plated on plastic alone (rather than poly-D-lysine) and maintained in GDM for 2 days.

Materials OL plated on (uncoated) plastic tissue culture dishes (from Basic Protocol 4, step 2; also see Fig. 2D.4.5) GDM medium (see recipe) Conditioned medium (from GDM; Basic Protocol 4)

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OLDEM medium (see recipe) Cell scraper Neuronal cultures (Support Protocol 4) plated on poly-D-lysine-coated coverslips Complete Neurobasal-N medium for cortical neurons (see recipe) 40-μm cell strainers (BD Falcon, cat. no. 352340) NOTE: All steps are performed at room temperature (20◦ C). 1. Detach OL cells with cell scraper and centrifuge at 450 × g, in the original culture medium. 2. Remove the supernatant and resuspend the cells in CM plus fresh OLDEM (1:2). See Figure 2D.4.5 (right side).

3. To prepare a single-cell suspension, which is necessary for the next step, remove any cell clusters by passing the cell suspension through a 40-μm sieve and wash the sieve as described in Basic Protocol 1, step 10. 4. Count the cells (UNIT 1C.3) and adjust the cell suspension to ∼200,000 cells/ml in OLDEM medium. 5. At 10 days after plating, remove half the volume (250 μl) of culture medium from neuronal culture (Support Protocol 4) without disturbing the cells. The neurons for coculture can be either cortical neurons or dorsal root ganglion cells.

6. Slowly add 200 μl of the OL suspension (from step 4) to the wells of one 24-well plate containing the neuronal cultures. To complete the original total volume in each well, add 50 μl/well of complete Neurobasal N medium for cortical neurons. 7. Follow the cocultures for at least 10 days. To feed, on day 5 after starting coculture, remove one half of the CM from each well (250 μl) and replace with fresh OLDEM. Repeat OLDEM feeding once a week. There is no need to reapply Neurobasal N medium. If the cultures are not overcrowded, they can be kept for at least 4 weeks. SUPPORT PROTOCOL 4

PREPARATION OF CORTICAL NEURONS Cortical neurons are one cell type that is used for coculture with the OLs to assess myelination.

Materials Complete Neurobasal-N medium for cortical neurons (see recipe) Poly-D-lysine-coated (see recipe) coverslips in wells of 12- or 24-well plates 37◦ C 4.5% CO2 incubator, 95% humidified Combustion Test Kit (Bacharach, cat. no. 10-500; http://www.bacharach-inc.com) Additional reagents and materials for isolation of rodent brain cells (see Basic Protocol 1) NOTE: All steps are performed at room temperature (20◦ C).


1. Prepare and dissociate embryonic rat brain tissue as described in Basic Protocol 1, steps 1 to 8, except use complete Neurobasal-N medium in step 5 of that protocol (instead of STMIIc) and dissect the brains in Neurobasal-N medium. 2. Add 2 to 4 ml of Neurobasal-N medium to the chunks left over in the dissociation tube and dissociate again five to eight times. 3. Filter the suspension of dissociated cells through 230-μm and 140-μm sieves to remove cell clusters.



4. Rinse the two sieves sequentially with Neurobasal-N containing 1% BSA at room temperature as described in Basic Protocol 1, step 10, and add this medium to the tubes containing the cells. 5. Collect the cells by centrifugation in the culture tubes 8 min at 400 × g. 6. Discard the supernatant, very gently as the pellet is very loose. 7. Resuspend the pellet in 4 ml of fresh Neurobasal-N medium with a 5-ml pipet by gently triturating (i.e., pipetting up and down) two or three times. Bring the volume to 12 ml (or the equivalent of 1 embryo/ml) with 2 parts of fresh Neurobasal N medium and 1 part of conditioned medium. 8. Assess cell viability with SYTOX (Support Protocol 2), count cells using a hemacytometer (UNIT 1C.3), and plate onto poly-D-lysine coated coverslips inside wells of 12or 24-well plates at 2 × 105 cells/well in 700 μl complete Neurobasal N medium for cortical neurons. 9. Incubate plated cells at 37◦ C with 4.5% CO2 /95% humidity and monitor CO2 with a Combustion Test Kit because most electronic panels do not give an accurate reading. 10. Every third day, add 50 μl/well complete Neurobasal-N complete medium. On the sixth day after plating, remove one-quarter of the medium and add the corresponding volume of fresh complete Neurobasal-N medium. Neurons are ready for coculture (Basic Protocol 5) 10 days after plating.

TRANSPLANTATION OF OL PROGENITORS INTO NEONATAL RATS Neural progenitor cells and their differentiated OL counterparts can be stereotaxically transplanted into the newborn developing rat brain relatively noninvasively as previously described (Snyder et al., 1997; Flax et al., 1998; Espinosa-Jeffrey et al., 2002). Similar results can be obtained with variations on the transplant method that are more suitable depending on the needs of the host brain and the type of study (Yandava et al., 1999; Ourednik et al., 2001, 2002; Park et al., 2002a; Teng et al., 2002; Wakeman et al., 2006; Lee et al., 2007; Redmond et al., 2007). A selection of detailed protocols for neonatal and adult mouse transplantation are described elsewhere (Espinosa de los Monteros et al., 1992, 1993a,b; Yan et al., 2004; Lee et al., 2008; Wakeman et al., 2009; UNIT 2D.3). Upon implantation into the lateral ventricles, donor cells engraft and migrate from the subventricular zone into the host corpus callosum, caudate putamen, and rostral migratory stream (RMS) in much the same manner as host NSC.

BASIC PROTOCOL 6

Materials Neonatal rat pup, post-natal day 0 to 5 (P0 to P5) 70% ethanol Dulbecco’s phosphate-buffered saline (DPBS; without calcium or magnesium, e.g., Cellgro, cat. no. 21-031-CV), sterile Microcentrifuge tube with cell sample (suspended in PBS; may be from various protocols in this unit depending on experimental question to be addressed) Borosilicate glass (Sutter Instrument Co., cat. no. B100–75-15) Micropipet puller (Sutter Instrument Co., Model P-87) Aspirator tube assemblies for calibrated microcapillary pipets (Sigma-Aldrich, cat. no. A5177–5EA) Fiber-optic light source for transillumination Warming pad Warm-water glove balloon

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Additional reagents and equipment for preparing injection micropipet (Lee et al., 2008) NOTE: Required materials may vary depending upon the grafting method of choice. 1. Prepare calibrated drawn borosilicate glass micropipet using borosilicate glass and a micropipet puller (Lee et al., 2008). 2. Anesthetize the neonatal rat pup by placing the pup on wet ice for ∼1.5 to 3 min until the animal no longer retains locomotion or responds to gentle toe and tail pinch. Carefully monitor the pup and immediately proceed to transplantation.

3. Insert a calibrated, drawn borosilicate glass micropipet into an aspirator tube assembly. Just prior to drawing up the cells, rinse the micropipet by drawing up and then expelling 5 μl of 70% ethanol five times, followed by sterile DPBS ten times to clean the needle.


Figure 2D.4.7 Human OL derived from (HFB-2050) human fetal NSC were labeled with fluorescent fast blue (FB; Sigma, cat. no. F-5756). A total of 60,000 cells were grafted into the corpus callosum (CC) of P(5) rat pups born to a myelin-deficient (md) carrier mother. At a time point 23 days after grafting, samples were harvested and examined. Grafted NSC survived and migrated extensively within the host brain parenchyma extending along the corpus callosum (CC) and caudate putamen (CPu). In the sketch, dots represent the location where FB+ cells were found. The sketch represents a sagittal view of the transplanted rat brain at 28 days of age, IS indicates where cells were implanted.



4. Gently flick sample in microcentrifuge tube prior to filling the needle, wipe the tube with 70% ethanol, and uncap the tube. 5. Slowly draw 4 to 5 μl cell suspension into the micropipet. 6. Loosely secure the head of the anesthetized pup and place directly over the light source to visualize the eyes and bregma. 7. Carefully insert the glass needle into the head at the midline between eye and bregma and slowly inject 2 to 5 μl cell suspension at 5 × 104 cells/μl into the lateral ventricle of either the left or the right hemisphere. Slowly remove the needle and check for leakage through the needle track. Repeat step 6 into the contralateral hemisphere. In addition to the lateral ventricles, NSC can also be transplanted into the striatum, the substantia nigra (SN), and corpus callosum (CC; Bjugstad et al., 2005, 2008; Redmond et al., 2007). Upon implantation into the CC of the host, HFB-2050 donor cells recognized by the fluorescent Fast Blue (FB) label migrated along the CC and into the caudate putamen (Fig. 2D.4.7). Precommitted OL can also be placed locally within focal sites of injury to decrease the need for extensive migration.

8. After the injection, warm the pup by placing on a warm-water glove balloon or heating pad to increase the body temperature before returning to the mother.


Basal stem cell medium (STM-II) Prepare 1 liter DMEM (low glucose, without glutamine, with sodium pyruvate; Invitrogen, cat. no. 11995-065). Supplement with the following:

5 mg insulin (Sigma, cat. no. I-5500) 50 mg transferrin (Sigma, cat. no. T-2252) 16.1 mg putrescine (Sigma, cat. no. P-7505) 20 nM (6.29 mg/liter) progesterone (Sigma, cat. no. P-7556) 8 μg sodium selenite (Sigma, cat no. S-5261); add 10 μl/liter of 0.8 mg/ml stock solution in PBS 2.2 g sodium bicarbonate (Fisher, cat. no. S233-500) 1 ml 10,000 U/ml penicillin/10 mg/ml streptomycin (Sigma, cat. no. P-4333) 1 ml 50 mg/ml kanamycin (Sigma, cat. no. K-0254) Store up to 2 weeks at 4◦ C This medium is used for rat and human NSC. STM-II is a variation of the original STM medium we previously described (Espinosa et al., 2002; UCLA case number 2002-475, formula available by Materials Transfer Agreement). STM-II yields results comparable to those obtained with STM.

Complete stem cell medium II (STMIIc) Just before use, combine 500 ml basal stem cell medium (STM-II; see recipe) and 500 ml NB-B27 medium (see recipe). This medium is used for plating, maintenance, and propagation of NSCs.

OL specification medium (OSM-II) Mix freshly prepared complete stem cell medium (STMIIc; see recipe) with freshly prepared glia defined medium (GDM; see recipe) at a 1:1 (v/v) ratio. This medium was formerly named OTM (Espinosa-Jeffrey et al., 2002); OSM-II derives from STM-II (see recipe).

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Glia defined medium (GDM) Combine 1 liter double-distilled water and one package DMEM/F12 medium (high glucose), then supplement with:

5 mg insulin (Sigma, cat. no. I-5500) 50 mg transferrin (Sigma, cat. no. T-2252) 16.1 mg putrescine (Sigma, cat. no. P-7505)) 2.2 g sodium bicarbonate (Fisher, cat no. S233-500) 4.6 g D-(+)-galactose (Sigma, cat. no. G-0625) 8 μg sodium selenite (Sigma, cat. no. S-5261): prepare 0.8 mg/ml stock solution in PBS (Sigma, cat. no. P-5368) and add 10 μl of this stock per liter medium 1 ml 50 mg/ml kanamycin (Sigma, cat. no. K-0254) Filter-sterilize through a 0.22-μm filter Prepare fresh From Espinosa de los Monteros et al. (1988, 1997)

Neurobasal-B-27 (NB-B-27) human neural stem cell proliferation medium Prepare 484 ml Neurobasal medium without Normocin, heparin, vitamin A, or LIF (Invitrogen, cat. no. 21103-049). Store up to 2 weeks at 4◦ C. Just before use in preparing STMIIc medium (see recipe), supplement with:

10 ml B-27 supplement without vitamin A (Invitrogen, cat. no. 12587-010) 5 ml GlutaMAX (Invitrogen, cat. no. 35050-061) 8 μg/ml heparin (Sigma, cat. no. H-3149) 2 ng/ml basic fibroblast growth factor (bFGF; Invitrogen, cat. no. 13256-029) 10 ng/ml leukemia inhibitory factor (LIF; Millipore, cat. no. LIF-1010) Neurobasal-N medium for cortical neurons, complete 1 liter Neurobasal medium (Invitrogen, cat. no. 21103-049) supplemented with: 5 mg insulin (Sigma, cat. no. I-5500)) 50 mg transferrin (Sigma, cat. no. T-2252) 8 μg sodium selenite (Sigma, cat. no. S-5261): prepare 0.8 mg/ml stock solution in PBS (Sigma, cat. no. P-5368) and add 10 μl of this stock per liter medium 2.2 g sodium bicarbonate (Fisher, cat. no. S233-500) 1 ml kanamycin (Sigma, cat. no. K-0254) Just before using, add the following to the supplemented Neurobasal-N to make the complete medium: 1:50 (v/v) B-27 supplement with vitamin A (Invitrogen, cat. no. 17504-044) 20 ng/ml recombinant basic bFGF (Invitrogen, cat. no. 13256-029) OLDEM medium Prepare glia defined medium (GDM; see recipe), but omit transferrin.

Poly-D-lysine coated wells/plates/coverslips


Prepare a stock solution by dissolving 100 mg poly-D-lysine in 100 ml water and filter sterilize through a 0.22-μm filter. Store in 5-ml aliquots at –20◦ C. When ready to use, dilute 1 part stock solution with 9 parts water to prepare 100 μg/ml working solution. Fill tissue culture dishes or wells with the working solution (and/or place coverslips to be coated into wells of 12- or 24-well plate) and incubate 1 hr in a humidified 37◦ C, 5% CO2 incubator, then remove solution by vacuum aspiration and allow surface to dry. Store coated tissue culture ware up to 3 months at 4◦ C. Use diluted solutions only once, but unused diluted aliquots can be stored up to 3 months at 4◦ C.



Tris-buffered saline (TBS) 2.42 g/liter Tris base 29.22 g/liter NaCl Adjust pH to 7.5 with HCl Store at room temperature up to 1 year, under sterile conditions COMMENTARY Background Information The described culturing system allows for the production of relatively homogeneous primary OL cultures in adequate numbers for cryopreservation. These cell stocks can be used for basic research in further in vitro studies. Moreover, these cells are never exposed to animal or human sera, and therefore remain as suitable candidates for cell replacement therapies in developmental disorders of the central nervous system (CNS) as well as neurodegenerative diseases. Numerous methods and culture media described in the literature (even before, the times of NSC) were the basis for the optimization of the culture media formulations described here (some examples are Botenstein and Sato, 1979; Saneto and de Vellis, 1985; Espinosa de los Monteros et al., 1988; Yang et al., 2005; UNIT 2D.1). Undoubtedly, all previous reports on how to obtain and culture OL derived from NSC have also been instrumental in designing the present protocols. For example, the group of Lachapelle and Baron-Van Evercooren described floating oligospheres derived from newborn rat brain (AvellanaAdalid et al., 1996). This concept has been applied to NSC to generate OLs by Zhang et al. (1998) and Espinosa-Jeffrey et al. (2002) and in the protocols in this unit. Zhang et al. (1998) described the use of B104 neuroblastoma cell–conditioned medium (B104CM) to induce the oligodendrocyte phenotype on neurospheres and induce proliferation. This approach provides OL for many kinds of studies, but they are unsuitable as donor cells for cell-replacement therapies to be used in translational studies, because they are produced using uncharacterized conditioned medium from B104 cells that have been grown in the presence of fetal bovine serum (as originally described by Louis et al., 1992). An example of the use of the protocols described can be found in Chattopadhyay et al., (2008).

Critical Parameters We want to emphasize that fate restriction towards commitment from NSC to OLP (as defined in Basic Protocol 3) becomes irreversible

after NSCs have been in OSM for at least 20 hr in either two- or three-dimensional cultures. Therefore, the progeny of these cells will define a homogeneous OLP population, ideal for biochemical, toxicological, and pharmacological studies, and also serve as an appropriate and reproducible source of committed cells to be used in cell therapy studies. Phenotype reversal of induced OLPs may be possible with genetic manipulation, but we have not attempted such studies to date. Always monitor the concentration of CO2 with a Combustion Test Kit, as most electronic panels do not provide an accurate reading. The proper lineage progression relies on precise control of CO2 to maintain a pH that should remain accurate and controlled.

Troubleshooting Human NSC are more fragile than their rodent counterparts; therefore, we recommend dissociation protocols that favor as little mechanical stress as possible. In our hands, enzymatic dissociation with 2 to 4 ml Accutase (Millipore) at 37◦ C for 3 to 5 min or light mechanical trituration through an 18-G needle (three to five times) is sufficient to dissociate hNSC into single cells and small (2- to 6-cell) clusters. Detailed methodology can be found elsewhere (Wakeman et al., 2009; UNIT 2D.3).

Anticipated Results OLPs obtained utilizing this system are plated on anti-PSA/NCAM plates and will attain a bipolar morphology if maintained in freshly supplemented OSM. Cells can also be plated directly onto plastic (tissue-culture grade). The morphology in that case may look more flattened or fibroblastic, but if maintained in fresh OSM, the early markers such as Olig2, Tf, PDGF-R, and NG2 will be expressed. At this stage, cells are still highly motile but will migrate less if plated onto poly-D-lysine. During this time, cells attain a more mature phenotype that truly represents their in vivo counterparts. Our culture media formulation includes the minimum and sufficient nutrients to support a given developmental stage; thus, cells cannot

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be kept indefinitely under these conditions because the substratum dictates the organization of the molecules on the cell membrane and poly-D-lysine confers a more permanent adhesion to the cells. Consequently, they would have the tendency to mature based on the signals coming from the cell membrane– substrate interaction (Linnemann and Bock, 1989; Mauro et al., 1994). Unfortunately, cells will not survive or remain healthy if maintained in unreplenished OSM as 2-D cultures, due to a lack of nutrients to support their transition to the next developmental stage. These cells will survive well if fed with OSM to renew the growth factors. The same concept applies to the transition to more mature OL stages. The nutrients and substrate together contribute to support cell signaling that will result in the formation of multiple cell processes followed by the synthesis of myelin components and their organization for membrane formation.

Time Considerations


The initial dissection and preparation of the primary cell suspension takes ∼2 hr. From the moment cells are plated on anti-PSA-NCAM (if fed regularly with fresh humoral factors), 100-mm dishes can be confluent within 3 to 4 days. Thus, generating 20 vials of rat NSCs for cryostorage would take ∼16 days. The generation of OLP from rNSC takes ∼24 hr; however, generating OLP in high numbers (15 vials) for storage would take 4 to 6 weeks. Lineage progression of rat OL towards more mature phenotypes takes ∼48 hr in the specific culture medium (GDM or OLDEM). In addition, OL will still proliferate in GDM, but at a much slower rate. Both GDM and OLDEM media are favorable to protein synthesis but less favorable for cell proliferation. Previously isolated ES cells and their NSC derivatives will need a longer period of time to provide high numbers of NSC for frozen stocks. This time will vary depending on the origin of the sample. We have had similar success directing NSC from several species, utilizing the same chemically defined media; however, incubation times may need to be increased for full maturation in higher-order mammals, such as primates. Induced cells lose NSC characteristics and acquire OLP features within 72 hr, yet their cell cycle is much slower, and, therefore, it would be necessary to propagate these cells 8 to 10 weeks to be able to create a healthy stock (six to eight vials) of human OLP. Previously established NSC lines (Snyder et al., 1992) can also be propagated

and specified into the OL phenotype using the system described here.

Acknowledgements A.E. and J. de V. thank the MRRC Media Core for preparation of figures and Dr. D. Birt for photograph of the dissection set up. This work (J.de V. and A.E.) was supported in part by PPGHD065-76 and by a Pilot grant from the National Multiple Sclerosis Society PP1498. D.R.W. thanks M. Hudson for critical review and comments. D.R.W. is supported in part by the American Parkinson’s Disease Association, HHMI Med-Into-Grad Training Fellowship, and the UCSD-NIH Training Fellowship in Clinical Genetics.

Conflict of Interest Statement The authors acknowledge no conflict of interest.

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protein in CaR-null mice. J. Neurosci. Res. 86:2159-2167. De Filippis, L., Lamorte, G., Snyder, E.Y., Malgaroli, A., and Vescovi, A.L. 2007. A novel, immortal, and multipotent human neural stem cell line generating functional neurons and oligodendrocytes. Stem Cells 25:2312-2321. Espinosa de los Monteros, A., Roussel, G, Neskovic, N.M. and Nussbaum, J.L. 1988. A chemically defined medium for the culture of mature oligodendrocytes. J. Neurosci. Res. 19:202-211. Espinosa de los Monteros, A., Zhang, M., Gordon, M., Aymie, M., and de Vellis, J. 1992. Transplantation of cultured premyelinating oligodendrocytes into normal and myelin-deficient rat brain. Dev. Neurosci. 14:98-104. Espinosa de los Monteros, A., Zhang, M.-S., and de Vellis, J. 1993a. O2A progenitor cells transplanted into the neonatal rat brain develop Into oligodendrocytes but not astrocytes. Proc. Natl. Acad. Sci. U.S.A. 90:50-54. Espinosa de los Monteros, A., Bernard, R., Tiller, B., Rouget, P., and de Vellis, J. 1993b. Grafting of fast blue labeled glial cells into neonatal rat brain: Differential survival and migration among cell types. Int. J. Dev. Neurosci. 11:625-639. Espinosa de los Monteros, A. and de Vellis, J. 1996. Vulnerability of oligodendrocytes to environmental insults: Potential for recovery. In The Role of Glia in Neurotoxicity (M. Aschner and H.K. Kimelberg, eds.) pp. 15-46. CRC Press, Boca Raton, Fla. Espinosa de los Monteros, A., Yuan, J., McCartney, D., Madrid, B.R., Cole, R., Kanfer, J.N., and de Vellis, J. 1997. Acceleration of the maturation of oligodendroblasts into oligodendrocytes and enhancement of their myelinogenic properties by a chemically defined medium. Dev. Neurosci. 19:297-311. Espinosa-Jeffrey, A., Becker-Catania, S., Zhao, P.M., Cole, R., and de Vellis, J. 2002. Phenotype specification and development of oligodendrocytes and neurons from rat stem cell cultures using two chemically defined media. J. Neurosci. Res. 69:810-825. Flax, J.D., Aurora, S., Yang, C., Simonin, C., Wills, A.M., Billinghurst, L.L., Jendoubi, M., Sidman, R.L., Wolfe, J.H., Kim, S.U., and Snyder, E.Y. 1998. Engraftable human neural stem cells respond to developmental cues, replace neurons, and express foreign genes. Nat. Biotechnol. 16:1033-1039. Kim, H.T., Kim, I.S., Lee, I.S., Lee, J.P., Snyder, E.Y., and Park, K.I. 2006. Human neurospheres derived from the fetal central nervous system are regionally and temporally specified but are not committed. Exp. Neurol. 199:222-235.

of immortal human neural stem cell line with multipotent differentiation property. In Methods in Molecular Biology, Vol. 438: Neural Stem Cells, 2nd ed. (L.P. Weiner, ed.) pp.103-121. Humana Press, Totowa, N.J. Larsen, E.C., Kondo, Y., Fahrenholtz, C.D., and Duncan, I.D. 2008. Generation of cultured oligodendrocyte progenitor cells from rat neonatal brains. Curr. Protoc. Stem Cell Biol. 6:2D.1.1-2D.1.13. Lee, H.J., Kim, K.S., Kim, E.J., Choi, H.B., Lee, K.H., Park, I.H., Ko, Y., Jeong, S.W., and Kim, S.U. 2007. Brain transplantation of immortalized human neural stem cells promotes functional recovery in mouse intracerebral hemorrhage stroke model. Stem Cells 25:12041212. Lee, J.P., McKercher, S., Müller, F.J., and Snyder, E.Y. 2008. Neural stem cell transplantation in mouse brain. Curr. Protoc. Neurosci. 42:3.10.13.10.23. Linnemann, D. and Bock, E. 1989. Cell adhesion molecules in neural development. Dev. Neurosci. 11:149-173. Louis, J.C., Magal, E., Muir, D., Manthorpe, M., and Varon, S. 1992. CG4, a new bipotential glial cell line from rat brain, is capable of differentiating in vitro either mature oligodendrocytes or type-2 astrocytes. J. Neurosci. Res. 31:193-204. Mauro, V.P., Wood, I.C., Krushel, C., Crossin, K.L., and Edelman, G.M. 1994. Cell adhesion alters gene transcription in chicken embryo brain cells and mouse embryonal carcinoma cells. Proc. Natl. Acad. Sci. U.S.A. 91:2868-2872. Müller, F.J., Snyder, E.Y., and Loring, J.F. 2006. Gene therapy: Can neural stem cells deliver? Nat. Rev. Neurosci. 7:75-84. Neman, J. and De Vellis, J., eds. 2008. Handbook of Neurochemistry and Molecular Neurobiology: Myelinating Cells in the Central Nervous System—Development, Aging, and Disease. Springer, New York. Ourednik, V., Ourednik, J., Flax, J.D., Zawada, W.M., Hutt, C., Yang, C., Park, K.I., Kim, S.U., Sidman, R.L., Freed, C.R., and Snyder, E.Y. 2001. Segregation of human neural stem cells in the developing primate forebrain. Science 293:1820-1824. Ourednik, J., Ourednik, V., Lynch, W.P., Schachner, M., and Snyder, E.Y. 2002. Neural stem cells display an inherent mechanism for rescuing dysfunctional neurons. Nat. Biotechnol. 20:11031110. Palmer, T.D., Schwartz, P.H., Taupin, P., Kaspar, B., Stein, S.A., and Gage, F.H. 2001. Cell culture: Progenitor cells from human brain after death. Nature 411:42-43.

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

Park, K.I., Teng, Y.D., and Snyder, E.Y. 2002a. The injured brain interacts reciprocally with neural stem cells supported by scaffolds to reconstitute lost tissue. Nat. Biotechnol. 20:1111-1117.

Kim, S.U., Nagai, A., Nakagawa, E., Choi, H.B., Bang, J.H., Lee, H.J., Lee, M.A., Lee, Y.B., and Park, I.H. 2008. Production and characterization

Park, K.I., Ourednik, J., Ourednik, V., Taylor, R.M., Aboody, K.S., Auguste, K.I., Lachyankar, M.B., Redmond, D.E., and Snyder, E.Y. 2002b. Global

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gene and cell replacement strategies via stem cells. Gene Ther. 9:613-624. Redmond, D.E. Jr., Bjugstad, K.B., Teng, Y.D., Ourednik, V., Ourednik, J., Wakeman, D.R., Parsons, X.H., Gonzalez, R., Blanchard, B.C., Kim, S.U., Gu, Z., Lipton, S.A., Markakis, E.A., Roth, R.H., Elsworth, J.D., Sladek, J.R. Jr., Sidman, R.L., and Snyder, E.Y. 2007. Behavioral improvement in a primate Parkinson’s model is associated with multiple homeostatic effects of human neural stem cells. Proc. Natl. Acad. Sci. U.S.A. 104:12175-12180. Saneto, R.P. and de Vellis, J. 1985. Characterization of cultured rat oligodendrocytes proliferating in a serum-free chemically defined medium. Proc. Natl. Acad. Sci. U.S.A. 82:3509-3513. Schwartz, P.H., Bryant, P.J., Fuja, T.J., Su, H., O’Dowd, D.K., and Klassen, H. 2003. Isolation and characterization of neural progenitor cells from post-mortem human cortex. J. Neurosci. Res. 74:838-851. Snyder, E.Y., Deitcher, D.L., Walsh, C., ArnoldAldea, S., Hartwieg, E.A., and Cepko, C.L. 1992. Multipotent neural cell lines can engraft and participate in development of mouse cerebellum. Cell 68:33-51. Snyder, E.Y., Yoon, C., Flax, J.D., and Macklis, J.D. 1997. Multipotent neural precursors can differentiate toward replacement of neurons undergoing targeted apoptotic degeneration in adult mouse neocortex. Proc. Natl. Acad. Sci. U.S.A. 94:11663-11668. Svendsen, C.N., Caldwell, M.A., and Ostenfeld, T. 1999. Human neural stem cells: Isolation, expansion and transplantation. Brain Pathol. 9:499-513. Teng, Y.D., Lavik, E.B., Qu, X., Park, K.I., Ourednik, J., Zurakowski, D., Langer, R., and Snyder, E.Y. 2002. Functional recovery following traumatic spinal cord injury mediated by a unique polymer scaffold seeded with neural stem cells. Proc. Natl. Acad. Sci. U.S.A. 99:3024-3029.

Villa, A., Snyder, E.Y., Vescovi, A., and Mart´ınezSerrano, A. 2000. Establishment and properties of a growth factor-dependent, perpetual neural stem cell line from the human CNS. Exp. Neurol. 161:67-84. Wakeman, D.R., Crain, A.C., and Snyder, E.Y. 2006. Large animal models are critical for rationally advancing regenerative therapies. Regenerative Med. 1:405-413. Wakeman, D.R., Hofmann, M.R., Teng, Y.D., and Snyder, E.Y. 2009. Derivation, expansion, and characterization of human fetal forebrain neural stem cells. In Human Cell Culture: Adult Stem Cells: Vol. 7. (J.R. Masters and B.O. Palsson, eds.). Springer, Dordrecht, The Netherlands. Williams, W.C. 2nd and Gard, A.L. 1997. In vitro death of jimpy oligodendrocytes: Correlation with onset of DM-20/PLP expression and resistance to oligodendrogliotrophic factors. J. Neurosci. Res. 50:177-189. Wysocki, L.J. and Sato, V.L. 1978. Panning for lymphocytes: A method for cell selection. Proc. Natl. Acad. Sci. U.S.A. 75:2844-2848. Yan, J., Welsh, A.M., Bora, S.H., Snyder, E.Y., and Koliatsos, V.E. 2004. Differentiation and tropic/trophic effects of exogenous neural precursors in the adult spinal cord. J. Comp. Neurol. 480:101-114. Yandava, B.D., Billinghurst, L.L., and Snyder, E.Y. 1999. Global cell replacement is feasible via neural stem cell transplantation: Evidence from the dysmyelinated shiverer mouse brain. Proc. Natl. Acad. Sci. U.S.A. 96:7029-7034. Yang, Z., Watanabe, M., and Nishiyama, A. 2005. Optimization of oligodendrocyte progenitor cell culture method for enhanced survival. J. Neurosci. Methods 149:50-56. Zhang, S.C., Lundberg, C., Lipitz, D., O’Connor, L.T., and Duncan, I.D. 1998. Generation of oligodendroglial progenitors from neural stem cells. J. Neurocytol. 27:475-489.




Isolation and Culture of Ventral Mesencephalic Precursor Cells and Dopaminergic Neurons from Rodent Brains

UNIT 2D.5

Jan Pruszak,1,2 Lothar Just,3 Ole Isacson,2 and Guido Nikkhah1 1

Freiburg University Hospital, Freiburg, Germany Harvard Medical School, McLean Hospital, Belmont, Massachusetts 3 Institute of Anatomy, Center for Regenerative Biology and Medicine, Eberhardt-Karls-University Tübingen, Tübingen, Germany 2

ABSTRACT The ability to isolate ventral midbrain (VM) precursor cells and neurons provides a powerful means to characterize their differentiation properties and to study their potential for restoring dopamine (DA) neurons degenerated in Parkinson’s disease (PD). Preparation and maintenance of DA VM in primary culture involves a number of critical steps to yield healthy cells and appropriate data. Here, we offer a detailed description of protocols to consistently prepare VM DA cultures from rat and mouse embryonic fetal-stage midbrain. We also present methods for organotypic culture of midbrain tissue, for differentiation as aggregate cultures, and for adherent culture systems of DA differentiation and maturation, followed by a synopsis of relevant analytical read-out options. Isolation and culture of rodent VM precursor cells and DA neurons can be exploited for studies of DA lineage development, of neuroprotection, and of cell therapeutic approaches in C 2009 by John animal models of PD. Curr. Protoc. Stem Cell Biol. 11:2D.5.1-2D.5.21. Wiley & Sons, Inc. Keywords: stem cells r cell and tissue culture r neuroscience r isolation r puriÞcation r separation r cell and developmental biology r cell therapy

INTRODUCTION This unit describes the dissection of the rat or mouse fetal ventral midbrain (VM) region (Basic Protocol 1) and the generation of dopamine (DA) neuronal cell cultures (Basic Protocols 2 and 3 and Alternate Protocols 1 and 2). Additionally, there is a description of analytical readouts (Support Protocol). These procedures have been applied previously in numerous in vitro and in vivo paradigms (Bjorklund et al., 1983; Nikkhah et al., 1994; Haque et al., 1997; Timmer et al., 2006). The detailed description and the synopsis of updated experimental concepts in this Þeld provided here may help promote a broader application and facilitate the study of midbrain DA neurons in the context of neural development and therapeutic models.

STRATEGIC PLANNING A ßow diagram of experimental options for study design is provided in Figure 2D.5.1. Due to the availability of numerous midbrain-relevant transgenic mice, mouse primary VM tissue can be considered advantageous for many experimental paradigms. This facilitates numerous functional neurobiological in vitro studies, and also comparative studies with murine pluripotent cell sources (Yurek and Fletcher-Turner, 2004; Chung et al., 2006; Lin and Isacson, 2006). Somatic Stem Cells Current Protocols in Stem Cell Biology 2D.5.1-2D.5.21 Published online December 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02d05s11 C 2009 John Wiley & Sons, Inc. Copyright


preparation of cell suspension

VM dissection

organotypic culture

aggregate culture

adherent culture

experimental read-out

Figure 2D.5.1 Overview: Isolation and culture of VM precursors and DA neurons. Dissection of rodent midbrain (see Basic Protocol 1) enables analysis of intact VM neural tissue for organotypic culture (see Basic Protocol 3), as well as gentle dissociation into single-cell suspensions (see Basic Protocol 2). Cell culture options include expansion and/or differentiation as three-dimensionalaggregate cultures (see Alternate Protocol 1), or as adherent monolayer cultures (see Alternate Protocol 2). Subsequent detailed analysis of DA neuronal phenotype is customized for the specific experimental paradigm at hand (see Support Protocol).

For transplantation studies, rat tissue has been more frequently applied, given the wide array of standardized behavioral tests in rat Parkinson’s Disease (PD) models (Dunnett, 1994). For immediate differentiation, resulting in maturing DA cultures within 1 to 3 days, use tissue from embryonic day 14 (E14) rats or E13 mice (DA neuronal culture). When a short-term expansion step is included (VM precursor culture), use tissue from E11.5 to E12 rats or E11 mice according to the subsequent protocols (see Alternate Protocol 2). Depending on the experimental question at hand, VM tissue of either stage can be used for Basic Protocols 2 or 3. NOTE: Experiments involving live animals must conform to national and institutional regulations and must be approved by the Institutional Animal Care and Use Committee (IACUC) or equivalent. Consider the scientiÞc and biomedical rationale for conducting the particular experiment. NOTE: Sterilize the instruments by autoclaving, and place in the aseptically prepared hood/clean dissection area. Ensure that assigned safety containers are available for scalpel blades and for the glass pipet waste. BASIC PROTOCOL 1

DISSECTION OF VENTRAL MESENCEPHALON This protocol is used to isolate the ventral mesencephalon (midbrain) from rodent embryos, embryonic day (E) 11 to E14, using microdissection techniques. Steps are described on how to identify anatomical landmarks, how to remove overlying tissue, and how to dissect the ventral midbrain portion itself. An accompanying video Þle (see Video 1 at http://www.currentprotocols.com/protocol/sc02d05) demonstrates the entire procedure.

Materials Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

C57B6/J mice (The Jackson Laboratory), embryonic day (E) 11 to E13 or Sprague–Dawley rats (Charles River), E11.5 to E14 Hanks balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170), ice cold Dissection buffer (see recipe)



Microdissecting instruments (sterilized; Fine Science Tools): Small dissecting scissors Medium dissecting scissors Dumont forceps—straight and angled or curved Curved microdissecting scissors Spatula Moria perforated spoon with holes Laminar ßow hood, sterilized by cleaning with 70% ethanol or UV-exposure for 15 min 60-mm and 100-mm round dishes (petri dishes), Þlled with dissection buffer Dissecting microscope (e.g., Leica MZ6 or Zeiss Stemi 2000) Curved scalpel blade (e.g., BD Bard-Parker no. 23 or 24) 15- and 50-ml conical tubes Collect embryos 1. Using aseptic technique, obtain the uterine horns from a time-pregnant mouse or rat [embryonic age E11.5-12 (rat) or E11.5 (mouse) for VM precursor cells; E14 (rat) or E13 (mouse) for direct DA neuron culture; see Strategic Planning and Fig. 2D.5.1]. 2. Submerge uterine horns in a 100-mm petri dish containing ice-cold, sterile CMFHBSS, and carefully rinse 2 to 3 times with 15 ml ice-cold, sterile CMF-HBSS. NOTE: From this point on work under sterile conditions in a laminar ßow hood, or add antibiotics (penicillin/streptomycin at standard concentrations) to reagents.

3. Transfer to a clean 100-mm petri dish containing dissection buffer. 4. Under a dissection microscope placed in a laminar ßow hood, perform the following steps (steps 5 to 19). Perform the steps in a timely manner, and keep the tissue cooled on ice and immersed in ice-cold buffers throughout the procedure. Follow the dissection sequence as depicted in Figure 2D.5.2. 5. Dissect each embryo from the uterine sac (Fig. 2D.5.2A-F), and remove the amniotic membranes (Fig. 2D.5.2G,H). 6. Using a Morian-type perforated spoon, transfer the embryo to a clean sterile petri dish containing ice-cold dissection buffer. 7. To conÞrm and monitor gestational age, measure and record crown rump length (CRL) of the embryos used. Expect CRL = 5 to 6 mm for early stage embryos (rat E11.5-E12; mouse E11; VM precursor culture). Expect CRL = 10 to 12 mm for later stage embryos (rat E14; mouse E13; DA neuronal culture).

8. Exclude any malformed or otherwise damaged embryos.

Dissect brains 9. Decapitate each fetus using microdissection scissors or a scalpel 10. Identify the central nervous system, and the midbrain region of interest (Fig. 2D.5.2J-M). CAUTION: The tip of the scissors should point away from the brain to avoid damaging the brain. See supplemental video material for a detailed demonstration of the procedure (see Video 1 at http://www.currentprotocols.com/protocol.sc02d05). Somatic Stem Cells


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A

B

C

D

E

F

G

H

I

J

K

L

M

N

O

Figure 2D.5.2 Dissection of the VM region from embryonic rodent brain. (A-F) Isolate the embryo from the uterine sac. (G-H) Free the embryo from any remaining placental and amniotic membranes. (I-L) Decapitate the embryo, and identify anatomical landmarks of the cranial central nervous system. Arrowheads indicate rostral and caudal borders of the midbrain region. Dotted lines outline the contour of forebrain CNS tissue. Arrow in (K) indicates VM region (lateral view). (L-M) Remove the overlying scalp tissue, to isolate the brain (superior view). Cut away the rostral forebrain and the caudal hindbrain regions (dashed lines; lateral view). (N) Open the resulting tube-like structure along the posterior midline (dashed line; coronal view). Arrow indicates anterior midline and VM region. (O) Trim the resulting butterfly-shaped structure, removing ∼2/3 of the posterior/lateral tissue on each side (dashed line; view from ventral midline, tissue flattened). Arrow indicates anterior midline of VM region. Also, see the supplemental Video 1 at http://www.currentprotocols.com/protocol/sc02d05. Abbreviations: fb, forebrain; hb, hindbrain; VM, ventral midbrain.

Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons



11. Carefully isolate the brain (Fig. 2D.5.2N). Holding the tissue with forceps near the forebrain or hindbrain regions (to be discarded in this VM DA cell isolation protocol), dissect away and remove the overlying scalp tissue. IMPORTANT NOTE: Avoid touching and damaging the midbrain region itself throughout the dissection procedure. The forebrain tissue can be processed in an identical manner, e.g., for comparative analysis or alternative studies.

12. Place the isolated brain in a clean 60-mm petri dish containing dissection buffer on ice. 13. Stabilizing the brain with forceps near the forebrain or hindbrain regions, carefully remove the fore- and hindbrain regions using a scalpel or microscissors. Place the rostral cut close to the forebrain vesicles and thalamic region and the caudal cut at the isthmus region (Fig. 2D.5.2M). If working with a scalpel, use the blade as a shield protecting the midbrain region, while removing the unwanted tissue with forceps (see Video 1 at http://www.currentprotocols.com/ protocol/sc02d05).

Dissect the ventral midbrain 14. Using forceps, steady the obtained midbrain tube (Fig. 2D.5.2O), exclusively touching the posterior midbrain region marked by the convex curvature at the dorsal midline. 15. Use small microscissors or the very tip of a curved scalpel blade to gradually dissect open this tube along the dorsal midline. 16. Carefully open the (Fig. 2D.5.2N,O).

now

characteristically

butterßy-shaped

tissue

ßap

17. Use forceps to thoroughly remove any remaining overlying meningeal tissue. Any remaining meningeal tissue can be recognized on the ventral exterior surface by its dense vascularization. When using the VM tissue for transplantation studies, such contaminating cells can promote unwanted immune reactions (Chen and Palmer, 2008). In vitro, the Þbroblast and endothelial cell types can overgrow and decrease the purity of the primary DA cultures.

18. Trim the outermost areas, i.e., the most dorsal parts of the midbrain tube, by dissecting away approximately two thirds of the tissue on each side (Fig. 2D.5.2O; i.e., approximately lateral/posterior to the sulcus limitans as an anatomical landmark). 19. Transfer the resulting tissue piece with dimensions of ∼0.3 × 1.0–mm into a conical tube containing cold dissection buffer kept on ice. Use ∼0.2 to 0.5 ml buffer volume per each VM. Note that, while immediate use is highly recommended, VM DA cell tissue pieces can in principle be stored up to 2 days at 4◦ C, in a hibernation medium and still yield viable VM DA cultures (Nikkhah et al., 1995). This strategy is utilized in clinical cell therapeutic paradigms, where human fetal VM DA tissue has been kept in hibernation medium supplemented with glial cell-derived neurotrophic factor (GDNF) prior to transplantation into patients (Mendez et al., 2005, 2008).

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BASIC PROTOCOL 2

PREPARATION OF CELL SUSPENSION This protocol is used for generation of a cell suspension from neural tissue (dissected in Basic Protocol 1). Such primary midbrain-derived cell preparations are used for in vitro culture or directly for transplantation assays. NOTE: For VM organ culture, skip this protocol and proceed directly as described in Basic Protocol 3. To obtain a cell suspension for use in three-dimensional aggregate cultures (Alternate Protocol 1) and/or adherent in vitro culture systems (Alternate Protocol 2), proceed as follows.

Materials Ventral midbrain tissue (Basic Protocol 1) Dissection medium (see recipe) Dissociation medium (see recipe) or trypsin 0.05% (w/v)/ EDTA (Invitrogen, cat. no. 25300) containing 0.2% (w/v) DNase I (see recipe) or Accutase (Innovative Cell Technologies, cat. no. AT104) or TrypLE Express (Invitrogen, cat. no. 12605) Heat-inactivated fetal bovine serum (FBS; Hyclone, cat. no. SH30070) Expansion medium (see recipe) Differentiation medium (see recipe) Trypan blue (Invitrogen, cat. no. 15250) or acridine orange/ethidium bromide solution (see recipe) 15-ml conical tubes Laminar ßow hood 37◦ C water bath Sterile Þre-polished 9-in. Pasteur pipets (see recipe) 200- and 1000-μl plastic tips and pipettors 70-μm cell strainer (BD, cat. no. 352350) or round bottom tube with 35-μm cell strainer caps (BD, cat. no. 352235) 1.5-ml microcentrifuge tubes Benchtop centrifuge Hemacytometer Microscope for viability dye detection (trypan blue: light microscope with bright Þeld or phase contrast; acridine orange/ethidium bromide: ßuorescence microscope with UV excitation and Þlters appropriate for simultaneous red-green channel detection; emission max for DNA is 526 nm, for RNA 650 nm) Additional reagents and equipment for determining the cell concentration and viability using trypan blue (UNIT 1C.3) Dissociate cells 1. Wash the obtained tissue pieces (Basic Protocol 1) in cold dissection buffer (e.g., 15 ml buffer in a 15-ml conical tube), by letting the tissue pieces sink down in the conical tube, pipetting off the medium, and Þlling the tube with fresh buffer. 2. Pipet off the buffer, and add 1 ml (per 10 midbrain tissue pieces) of the solution to be used for dissociation. Use either dissociation medium, or alternatively 0.05% trypsin/EDTA with 0.2% (w/v) DNase added, Accutase, or TrypLE Express. Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

To reduce the clumping of cell suspensions due to sticky nucleic acids released by damaged cells during dissociation, the addition of DNase during mechanical dissection and when using trypsin/EDTA or TrypLE Express is highly recommended (Panchision et al., 2007; Pruszak et al., 2007). The commercially available preparation Accutase already contains DNase activity.



3. For enzymatic digestion, incubate 3 to 15 min at 37◦ C. Use visual control and determine the optimal duration by test trituration. Avoid over-digestion, and inactivate with 10% fetal bovine serum if required (for trypsin/EDTA digestion). This is a critical step. Obtaining a viable neural cell suspension is a prerequisite for the subsequent culture protocols.

4. Using Þre-polished Pasteur pipets with decreasing diameter, gently dissociate the tissue pieces, ∼20 times total. Alternatively, trituration may be performed using Þrst a 1000-μl pipettor, followed by trituration with a 200-μl pipettor. Avoid excessive formation of air bubbles during mechanical dissociation of VM tissue, as those will reduce cell viability.

5. In case major tissue chunks remain in the solution, selectively triturate those pieces separately. Consider discarding some tissue, rather than compromising the major part of the cell suspension due to mechanical dissociation. For an optional Þltering step, pipet the obtained cell suspension through a cell strainer cap or through a 35- to 70-μm mesh. To minimize loss of cells due to this step, subsequently ßush the Þlter membrane with a small volume of medium.

6. Centrifuge 3 to 5 min at 200 × g, 4◦ C . Pipet off supernatant. 7. Resuspend in plating medium (either expansion medium or differentiation medium). The amount of plating medium varies (e.g., 200 μl per 10 midbrain pieces originally isolated).

Determine cell number and viability 8. Determine the cell concentration and the viability of an aliquot of the cell suspension, using a classic dye exclusion method (trypan blue; UNIT 1C.3) or DNA/RNA labeling techniques (acridine orange/ethidium bromide). Use a small sample of the obtained cell suspension and dilute it with the viability test dye, at a ratio of 1:10 (e.g., 1 or 5 μl cell suspension and 9 or 45 μl of dye, respectively). 9. After gentle mixing, transfer the sample solution with a fresh pipet tip to a hemacytometer chamber for visual inspection and quantiÞcation under a microscope. 10. Determine the viability and calculate the cell concentration. In trypan blue staining, dead cells will take up the dye and will appear blue under a light microscope. With acridine orange/ethidium bromide, live cells appear green (acridine orange due to labeling of RNA), while dead cells are labeled red (due to acridine orange and ethidium bromide labeling of DNA) upon UV-excitation and detection in the 520 to 650 nm range on a ßuorescence microscope. Cell viability needs to be higher than 80%, and should routinely range from 95% to 100% (see Fig. 2D.5.4A). Note that DA neurons are among the most fragile cells in the solution, and while cultures will contain neuronal cell types after relatively harsh treatment, the DA numbers will be low. Use the live cells counted for calculating the cell concentration.

11. Keep the cell suspension on ice or at 4◦ C until use. Before proceeding with either the three-dimensional aggregate (Alternate Protocol 1) or the adherent culture steps (Alternate Protocol 2), or with transplantation experiments, consider including puriÞcation methodologies recently optimized for neural cell suspensions such as ßuorescence-activated cell sorting (FACS) or immunomagnetic cell separation (MACS; Kerr et al., 1994; Ono et al., 2007; Panchision et al., 2007; Pruszak et al., 2007); also see the ßow chart in Fig. 2D.5.1).

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BASIC PROTOCOL 3

MIDBRAIN NEURAL CULTURE: ORGANOTYPIC CULTURE This protocol outlines the detailed procedures for in vitro culture of VM precursor and DA neuronal cell types. One option is to culture midbrain tissue obtained in Basic Protocol 1 as intact organotypic cultures on a membranous insert in the well. This preserves, to some extent, a physiological cellular context. An alternative option is the formation of three-dimensional aggregate cultures. Sectioning such spherical aggregates provides a good readout of cells grown in near-physiological conditions (Alternate Protocol 1). The most commonly applied option is the culture of VM neural precursors or DA neurons on permissive substrates such as laminin and/or poly-L-ornithine (Alternate Protocol 2). NOTE: Either early stage (VM precursors, E11.5-12 rats, E11 mice) or later stage fetal tissue (DA neurons, E14 rats, E13 mice ) can be used. However, the efÞciency for a short-term expansion step is limited to the early stage VM precursors (E11.5-12 rat; E11 mouse; CRL ≈5 to 6 mm. See Alternate Protocol 2).

Materials Ventral midbrain tissue pieces (Basic Protocol 1) Differentiation medium (see recipe) 4% (w/v) paraformaldehyde (PFA) solution Laminar ßow hood Pasteur pipet with a Þre-polished widened oriÞce (see recipe) or curved forceps Forceps or tungsten needles Millicell cell culture inserts (for six-wells; e.g., Millipore, cat. no. PICM0RG50) 6- and 24-well tissue culture plates (e.g., Fisher, Falcon or Nunc) 37◦ C water bath Scalpel Perform organotypic cultures 1. After isolating the midbrain tissue (characteristic “butterßy” structure, Basic Protocol 1; see Video 1 at http://www.currentprotocols.com/protocol/sc02d05; see Fig. 2D.5.2O), carefully transfer the intact VM tissue piece to a membrane-covered tissue culture insert, either by using a Pasteur pipet with a Þre-polished widened oriÞce, or by carrying the tissue in a liquid droplet between the tips of the branches of a curved pair of forceps (Fig. 2D.5.3A). 2. Using forceps or tungsten needles, gently ßatten the tissue and orient it towards the central area of the insert, away from the edges. 3. Using a pipet, very carefully remove any remaining ßuid on the top surface of the insert’s membrane, to ensure adherence to the membrane and to avoid ßoating of the tissue. This is a critical step to avoid ßoating of the tissue.

4. Place the insert into the well of a six-well culture plate Þlled with 1.5 ml differentiation medium. Ensure that no air bubbles remain below the insert’s membrane. 5. Culture for up to 10 days, while changing medium every two days in the compartment below the insert.



6. For staining and analysis of this in vitro system, Þx the tissue in the well 20 min at room temperature with 4% paraformaldehyde solution, and carefully cut out the membrane from the insert using a scalpel blade. 7. Transfer the membrane piece with the VM tissue attached to a smaller well format (4-well or 24-well) for immunocytochemistry. 8. For further speciÞcs regarding staining and analysis, refer to Support Protocol. Current Protocols in Stem Cell Biology

A

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TH

E12

C

TH

D

TH

Figure 2D.5.3 VM region organotypic culture. (A) Transfer of intact VM tissue onto tissue culture insert. (B) Precursor stage (rat E12) VM region organ culture stained for the DA marker TH. (C) Rat E12 VM region organotypic culture after 2 days in vitro, stained for TH. (D) Higher magnification of VM DA neurons in intact VM region tissue culture. Abbreviations: TH, tyrosine hydroxylase; DA, dopaminergic; VM, ventral midbrain; E12, embryonic day 12; div, days in vitro. Scale bars = 100 μm.

MIDBRAIN NEURAL CULTURE: THREE-DIMENSIONAL AGGREGATE CULTURE


An alternative for culturing midbrain neural cells after dissociation is the three dimensional neural culture (see Fig. 2D.5.4).

Additional Materials (also see Basic Protocols 2 and 3) VM cell suspension (Basic Protocol 2) Differentiation or expansion medium (see recipes) 4% (w/v) paraformaldehyde solution 15% (w/v) agar gel 15-ml conical tubes Shaker/roller tube system (e.g., Miltenyi Biotec, cat. no. 130-090-753, MACSmix Tube Rotator) HumidiÞed tissue culture incubator (37◦ C, 5% CO2 ), preferably including low O2 option Vibratome 1. Dilute the obtained cell suspension appropriately. For aggregate culture, transfer ∼1–5 × 105 live VM cells dissolved in 5 ml medium into a 15-ml conical tube. Somatic Stem Cells


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A

AO/EthBr

B

E12

C

TuJ1/DAPI

D

TH

Figure 2D.5.4 Three-dimensional-aggregate culture. (A) Viability of VM cell suspensions for three-dimensional-aggregate cultures, adherent culture systems, and transplantation studies alike is determined by viability dyes. Here, acridine orange/ethidium bromide (fluorescent image showing live cells in green; upper panel). Lower panel: phase contrast image of the identical field. (B) Aggregate formed after 7 days in vitro in the roller tube system. (C) Aggregate cultures stained for nuclear marker DAPI (tightly packed), TuJ1 neuronal marker, dense fiber network surrounding. (D) Aggregate cultures stained for tyrosine hydroxylase. Inset: higher magnification. Sections of aggregates cut on a vibratome after embedding in agarose are shown; C and D display identical areas. Abbreviations: AO, acridine orange; EthBr, ethidium bromide; TuJ1, neuronal marker beta-III tubulin; DAPI, nuclear marker; TH, tyrosine hydroxylase.

2. Place the tube into a shaker/roller tube system that enables a steady rocking or turning motion of about 30 cycles per min, inside a tissue culture incubator. NOTE: Set up the roller tube system (Alternate Protocol 1) inside the tissue culture incubator. Arrange in such a way that liquid in the tubes will not leak out (if needed, position the system in a tilted but stable manner).

3. Culture for 3 to 10 days. Change medium every other day, by letting the cells settle/sink to the bottom, placing the tube vertically for ∼10 min, and then carefully removing the supernatant, leaving a small volume of medium to avoid cell loss. After 3 to 5 days, macroscopically visible clusters of cells will have formed.

4. For staining and analysis of this in vitro system, Þx the cells in the tube with 4% paraformaldehyde solution. Embed each aggregate in a 15% agar gel. Isolation and Culture of Ventral Mesencephalic Precursors Cells and Dopaminergic Neurons

5. Carefully cut each aggregate using a vibratome into sections of ∼20-μm thickness. Those sections can be further processed according to standard procedures such as immunocytochemistry. For further speciÞcs regarding staining and analysis, refer to Support Protocol.



MIDBRAIN NEURAL CULTURE: ADHESION CULTURE Adhesion cultures of VM cells enable the three-dimensional culture of dissociated cell suspensions (see Fig. 2D.5.5).


Materials VM cell suspension (Basic Protocol 2) Expansion medium (see recipe) Differentiation medium (see recipe) 24-well tissue culture plates Laminin/poly-L-ornithine coated 12-mm coverslips (see recipe) 100- or 200-μl pipets HumidiÞed tissue culture incubator (37◦ C, 5% CO2 ), preferably including low O2 option Dilute the obtained cell suspension 1a. For expansion of VM precursor cells (E11.5-E12 rats; E11 mice): Plate the cells in 24-well tissue culture plates at densities of 1–5 × 104 cells per cm2 in 0.5 to 1.5 ml expansion medium including 20 ng/ml bFGF as a mitogen. 1b. For differentiation of DA neuronal cell suspension (E14 rats; E13 mice): Plate at densities of 2–5 × 105 cells per cm2 in 0.5 to 1.5 ml differentiation medium. 2. To maximize the yield, plate cells as 30-μl droplet cultures, using a 100-μl or 200-μl pipet, on a freshly coated, washed, and brießy air-dried coverslips. A 30-μl drop covers an area of ∼1 cm2 .

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B

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Nestin/TuJ1/DAPI

D BrdU/total [%]

50

0

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TuJ1/DAPI

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TiJ1

2 div

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Figure 2D.5.5 Adherent culture system. (A) VM DA precursors (E11-12) at 1 day in vitro after plating, and (B) at 7 days in vitro of expansion with bFGF. (C) VM DA precursors during the expansion phase stain positive for Nestin (red), a minor fraction of cells stains positive for beta-III-tubulin (green). Blue = nuclear Hoechst stain. (D) Expansion and proliferative capacity is monitored by BrdU incorporation assays: here ranging from 44.2% BrdU+ cells at 1 day in vitro, to over 46.4% at 5 days in vitro to 30.8% at 7 days in vitro. Error bars indicate SEM; three independent experiments. (E) Differentiation of DA neurons is induced subsequent to in vitro expansion or alternatively immediately after VM dissection from older embryos (E14), forming a dense network of neuronal processes, staining positive for neuronal markers such as beta-III-tubulin (TuJ1, green; F, G) and dopaminergic markers such as tyrosine hydroxylase (TH, red; H).


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Calculate the cell concentration of the cell suspension accordingly. For example, for plating of 20 coverslips of E11.5 VM precursor cells as 30-μl droplets (area ≈1-cm2 ), at a density of 20,000 cells per cm2 , 400,000 cells suspended in 600 μl expansion medium are needed. Consider loss due to pipetting and measuring errors, and prepare cell suspension in surplus.

3. After gentle mixing of the cell suspension (avoid additional trituration), position a 30-μl cell suspension droplet onto the center of a coverslip in a 24-well plate. 4. Repeat until all wells are plated with the available cell suspension in the same manner. 5. Using caution not to spill the droplets, place the tissue culture plate into an incubator for ∼0.5 hr. 6. Carefully Þll up the wells to a volume of 0.5 to 1.3 ml with expansion or differentiation medium per 24-well. When too much debris is present, a gentle washing step, adding fresh medium, can be performed at this stage. For VM precursor cultures use expansion medium, for DA neuronal cultures use differentiation medium. Ensure that the coverslips are entirely covered by medium. Remove potential air bubbles under the coverslips, as those may later cause ßoating of the coverslips, and drying of the surface area.

7. For the VM expansion cultures, change medium from expansion medium to differentiation medium after 2 to 4 days. 8. Culture for 3 to 10 days, changing the medium every other day. Depending on the cell density and resulting metabolic turnover, daily medium changes should be considered at prolonged stages in culture (monitor pH/phenol red indicator property of medium). DA neuronal primary cultures are very sensitive. Perform the medium changes rapidly and carefully, to avoid letting any well dry.

9. Process for analysis (see Support Protocol). SUPPORT PROTOCOL

ANALYSIS OF VM NEURAL PRECURSORS AND DA NEURONS This section summarizes and discusses the analytical readout options available to study VM neural precursors and DA neurons (see Fig. 2D.5.6).

Materials


Dulbecco’s phosphate-buffered saline (DPBS) Mg+, Ca+-free (CMF-DPBS; Invitrogen, cat. no. 14190) Antibodies typically used in a basic VM DA differentiation: Sheep anti-TH (1:1,000; Pel-Freez) Mouse anti-nestin (1:100; Millipore/Chemicon) Rabbit anti-TuJ1 (Covance 1:1000) Mouse anti-MAP2 (Millipore/Chemicon 1:500) Mouse anti-Pitx3 (Zymed 1:1000) Rabbit anti-Pitx3 (1:250; Invitrogen) Rabbit anti-glial Þbrillary acidic protein (1:500; Dako) Rabbit anti-Nurr1 (E-20; 1:300; Santa Cruz Biotechnology) Mouse anti-engrailed 1 (clone 4G11; 1:40) Rabbit anti-ki67 (1:2,000; Novocastra/Vector Laboratories) Rabbit anti-DAT (1:1000; Millipore/Chemicon) Corresponding secondary antibodies Pipets



Microscope for cell analysis Vibratome for sectioning of three-dimensional-aggregate cultures (Leica VT1000 S; Basic Protocol 3) Analysis of midbrain neural precursor and DA neuronal cultures Staining protocols include immunohistochemical, e.g., immunoperoxidase-based, and immunoßuorescence techniques. For a detailed description of these procedures, refer for example, to Glynn and McAllister, 2006; Kim et al., 2006; or Hoffman et al., 2008. Commonly used antibodies for a basic analysis of neural differentiation include those against the neural precursor marker nestin, and neuronal markers such as TuJ1 (beta-3tubulin), MAP2, or Tau. Proliferation assays may be performed using bromodeoxyuridine (BrdU) incorporation to determine the expansion potential of neural precursors. Astroglial differentiation can be documented by staining for glial Þbrillary acidic protein (GFAP). Basic analysis of midbrain DA phenotype includes staining for the catecholaminergic marker tyrosine hydroxylase (TH), co-labeled with DA transcription factors such as Pitx3 and/ or Nurr1. The A9 DA neurons speciÞcally express the marker GIRK2. Stained neurons in vitro may be analyzed with respect to neurite outgrowth, directed targeting, or the inßuence of co-culture conditions, e.g., with mesencephalic glial cell types. Supplementing such immunoßuorescence and morphological studies, phenotypic characterization is also done by gene expression analysis via RT-PCR, and by protein analysis (Western blot; Sonntag et al., 2007). SpeciÞc subpopulations can be analyzed by using techniques such as laser capture microdissection (Espina et al., 2006), or ßow cytometric analysis of cell subsets labeled with antibodies or transgenic ßuorescent markers (Pruszak et al., 2007).

A

TuJ1/Hoechst

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TuJ1/GFAP/Hoe

post-LCM

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post-FACS

1000 CD24HIGH

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Figure 2D.5.6 Options for analytical readout of VM DA neurons. Immunocytochemical assays include measures of neurite outgrowth and targeting studies (A), and/ or co-culture assays of VM DA neurons, here using astroglial feeder cells (B). Detailed analysis of specific DA neuronal subsets is achieved by isolating fixed DA neurons using laser capture microdissection (LCM) (C). Fluorescence-activated cell sorting (FACS) methods optimized for fragile neural cell types enables isolation of viable VM DA neurons for further in vitro and in vivo analysis in pharmacological, toxicological, and cell transplantation assays (D,E).


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Functional aspects of live VM DA neurons in vitro may be studied through electrophysiological analysis (Schlesinger et al., 2004). Furthermore, HPLC serves to detect dopamine release and dopamine metabolites such as DOPAC in media supernatants and/or cell samples as additional parameters for evaluating the extent of DA differentiation and the functionality of the cultured cells. Typically, the latter test is performed under basal conditions and then after stimulation, e.g., with 56 mM potassium chloride solution. See Studer et al. (1998) and Chung et al. (2002) for details. Finally, in vivo transplantation of VM DA neurons, commonly ectopically into the striatum of rodent animal models, enables the study of integration of VM DA neurons into the host circuitry by histological, as well as behavioral analyses (Vinuela et al., 2008; Nikkhah et al., 2009).


Acridine orange/ethidium bromide solution Prepare a solution of 5 μg/ml acridine orange (Sigma, cat. no. A8097) and 5 μg/ml ethidium bromide (Sigma, cat. no. E151) in phosphate-buffered saline (e.g., CMF-DPBS; Invitrogen, cat. no. 14190). Store up to 12 months at 4◦ C. CAUTION: Acridine orange/ethidium bromide solution is mutagenic and is a health hazard. This solution (10-fold) can be prepared and stored at 4◦ C, protected from light, for 3 months).

Ascorbic acid (AA) stock solution (1000×) Dissolve ascorbic acid (Sigma, cat. no. A4034) in phosphate-buffered saline (CMFDPBS; Invitrogen, cat. no. 14190) to prepare a 200 mM stock solution, and Þltersterilize using a 0.22-μm Þlter. Protect from light. Store up to 12 months at −20◦ C. Basic Þbroblast growth factor (bFGF) stock solution Dissolve bFGF (Invitrogen, cat. no. PMG0034) in phosphate-buffered saline (CMFDPBS; Invitrogen, cat. no. 14190) at 2 μg/ml. Divide into 50-μl aliquots and store up to 6 months at −20◦ C. Differentiation medium Neurobasal medium (Invitrogen, cat. no. 21103) L-glutamine (Invitrogen, cat. no. 21051-016) 50× B27 (Invitrogen, cat. no. 17504) 1% (v/v) heat-inactivated fetal bovine serum (FBS; Hyclone, cat. no. SH30070) Add 100 μM ascorbic acid (see recipe) immediately before use Store up to 7 days at 4◦ C ModiÞcations known to enhance the DA fraction include the addition of growth factors (see recipe) such as GDNF (Costantini and Isacson, 2000), FGF20 (Ohmachi et al., 2003), or TGF3beta to the differentiation medium.



Dissection buffer Hanks balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170) 20 mM D-glucose (Sigma, cat. no. G8270) Penicillin/streptomycin (Invitrogen, cat. no. 15140; use standard concentrations, as indicated by the supplier) Just before use, add 100 μM ascorbic acid (see recipe) Shelf life: 1 week at 4◦ C Current Protocols in Stem Cell Biology

Dissociation medium DMEM/F12 (Invitrogen, cat. no. 11320) 0.05% (w/v) DNase (Sigma, cat. no. D5025) 50× B27 (Invitrogen, cat. no. 17504) Prepare fresh DNase I stock Dissolve DNase I (Sigma, cat. no. D5025) at 0.2 mg/ml in Hanks’ balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170). Divide into aliquots and store up to 12 months at −20◦ C. Prepare sufÞcient quantity by dissolving 0.0024 g of DNase in 12 ml of HBSS/glucose (see recipe; this large volume allows for several rinses and trituration). Sterile Þlter using a 0.22-μm Þlter and keep on ice; then use or freeze.

Expansion medium DMEM/F12 (Invitrogen, cat. no. 11320) 100× N2 (Stem Cell Technologies, cat. no. 07152) Add 20 ng/ml bFGF (see recipe) immediately before use Store up to 2 weeks at 4◦ C Refer to Bouvier and Mytilineou (1995) and Studer et al. (1998) (the original papers introducing bFGF supplementation in the expansion medium).

Fire-polished glass pipets Prepare a set of Þve to ten Þre-polished pipets over a small ßame (alcohol or natural gas burner), such that the edges are smoothed and with decreasing aperture diameter starting at ∼1 mm (Schnitzler et al., 2008). Autoclave.

Growth factors Prepare as 1000× stocks and divide into aliquots. Store up to 6 months at −80◦ C. Add to the expansion or differentiation medium (see recipes) just before plating or medium change.

HBSS/glucose solution Dissolve 1.08 g D-glucose (Sigma, cat. no. G8270) in 500 ml Hanks’ balanced salt solution Mg2+ , Ca2+ -free (CMF-HBSS; Invitrogen, cat. no. 14170) to prepare a 20 mM glucose solution and Þlter-sterilize using a 0.22-μm Þlter. Store up to 2 weeks at 4◦ C.

Laminin solution Thaw laminin stock (Becton-Dickinson, cat. no. 354232) slowly to avoid gelatinization. Prepare on ice, using cooled pipets (kept at −20◦ C) to a Þnal concentration of 1 μg/ml in CMF-DPBS. Use immediately. CAUTION: Laminin rapidly adheres to surfaces (i.e., vessels, pipets, tips) if not kept ice-cold throughout preparation.

Poly-L-ornithine 0.01% (w/v) solution (Sigma, cat. no. P4957) Þnal. Dilute stock solution of polyL-ornithine [5 mg/ml in Dulbecco’s phosphate-buffered saline (DPBS) Mg++ , Ca++ -free (CMF-DPBS; Invitrogen, cat. no. 14190] and Þlter-sterilize using a 0.22-μm Þlter. Store up to 2 weeks at 4◦ C.

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Poly-L-ornithine/laminin-coated plates Coat plates (six-wells or 24-wells with glass coverslips) sequentially with poly-Lornithine (15 μg/ml; see recipe) and then laminin (1 μg/ml; see recipe) solutions. Throughout, ensure to add sufÞcient quantity of coating solutions or washing buffer to cover the surface of culture plates. First, incubate with poly-L-ornithine solution (15 μg/ml; see recipe) for 2 hr at 37◦ C or overnight at room temperature. Remove solution, wash with deionized water. Second, add freshly prepared ice-cold laminin solution (1 μg/ml; see recipe) using cooled pipets, and incubate for 2 hr at 37◦ C or overnight at room temperature. Remove solution, wash three times with 2 ml deionized water. In our hands, we obtain most consistent results using the coated plates right away, either “wet” or after brief (15 min) air drying in the hood. Note that dry coverslips are required for droplet culture (see Alternate Protocol 2).



Dopamine (DA) neurons of the substantia nigra region (pars compacta), A9 region, in the ventral midbrain (VM) play a critical role in the initiation of movement, as well as in behavior, motivation, emotion, and cognition through their contribution to basal ganglia neural circuitry (Ungerstedt, 1976; Fibiger and Phillips, 1988; Koob and Swerdlow, 1988; Perrone-Capano and di Porzio, 1996). Their progressive degeneration is a major feature of Parkinson’s disease (Olanow, 2007), leading to signiÞcant motor disability (Weintraub et al., 2008). DA neurons are generated from midbrain (mesencephalic) precursor cells, which originate from the neuroepithelial layer in the ventral neural tube of that region (Prakash and Wurst, 2006; Smits et al., 2006; Ono et al., 2007; Smidt and Burbach, 2007). Transcription factors relevant for the in vivo speciÞcation of DA neurons have been identiÞed in signiÞcant detail utilizing wild-type and DA-relevant transgenic mouse strains, such as Pitx3 (Smidt et al., 2004a,b), Lmx1b (Asbreuk et al., 2002), Otx2 (Puelles et al., 2004; Borgkvist et al., 2006), Nurr1 (Saucedo-Cardenas et al., 1998; Smits et al., 2003), Fox2a (Ferri et al., 2007), and Ngn2 (Andersson et al., 2006; Thompson et al., 2006). Functional DA neurons have also been generated from embryonic stem cells (Lee et al., 2000; Barberi et al., 2003; Perrier et al., 2004; Chung et al., 2006), and more recently mouse induced pluripotent stem (iPS) cells (Wernig et al., 2008). In the induction of DA neurons from such pluripotent cell sources, a major fraction has been shown to express DA-speciÞc markers, and the proofof-principle of restoring function in PD animal models has been demonstrated (Bjorklund

et al., 2002; Sonntag et al., 2007; Wernig et al., 2008). However, such DA patterning efforts from pluripotent sources require further optimization and Þne-tuning (Pruszak and Isacson, 2009), as so far only a smaller fraction of DA neurons derived from pluripotent cell sources may actually be patterned toward a true equivalent of the physiological phenotype (Hedlund et al., 2008). Thus, DA cells obtained from primary VM tissue represent a physiologically derived gold standard for studies of DA neuronal features, such as lineage development (Thompson et al., 2006; Ono et al., 2007), selective vulnerability (Chung et al., 2005, 2007), and functionality in behavioral (Dunnett, 1994; Klein et al., 2007) and electrophysiological assays (Geracitano et al., 2005; Rick et al., 2006). They provide a means for pharmacological and toxicity testing (SalthunLassalle et al., 2004; Chung et al., 2005), and serve as a valuable reference point for the derivation of DA neurons from pluripotent stem cells (Lin and Isacson, 2006).

Critical Parameters and Troubleshooting Culturing DA neurons Low yield of DA neurons is often a result of (1) too generous dissection of VM region, (2) too harsh trituration of VM tissue or (3) too long an exposure to enzymatic digestion during the cell preparation process. ModiÞcations known to enhance the DA fraction include the addition of GDNF (Costantini and Isacson, 2000), FGF20 (Ohmachi et al., 2003), or TGF3beta to the differentiation medium. Viability Similarly, low overall viability is usually due to harsh trituration generating many air



bubbles, or extended enzymatic digestion during the cell preparation process. Use a Þrepolished Pasteur pipet with gentle trituration. Adherence Poor adherence after plating suggests that the quality of the coating is not sufÞcient to induce or support DA neuronal growth. Prepare coating solutions freshly, keep them cool, and apply them promptly. If necessary, precoating of poly-L-ornithine/laminin-coated dishes with 10% FBS in PBS for ∼10 min prior to plating can improve adherence (remove FBS solution from the well before plating cells). Gently wash and change to fresh medium the following day, particularly for the VM expansion protocol, as serum components will induce differentiation. Plating density Higher-density plating (e.g., 0.5 × 106 cells/cm2 ) will often yield better quality cultures with improved appearance; however, analysis on a single-cell level can be difÞcult in such dense cultures. Note that more frequent medium changes and increasing the medium volume per well will be required for culture plated at higher densities. Contamination Under aseptic conditions, VM DA cultures can be grown without antibiotics, which is the preferred method. Antibiotics can foster bacterial resistance and could interfere with physiological tests performed. Nevertheless, penicillin-streptomycin is still widely applied in tissue culture as a preventative measure. Other suspected contamination of VM DA cultures can be successfully treated with additional supplementation of Normocin (Invivogen, cat. no. ant-nr-o) to the medium. Mycoplasma spp. are generally less of an issue in primary culture, but cross-contamination from existing cell-line work in the laboratory cannot be excluded. After positive testing of cell cultures for Mycoplasma spp., eradication can be attempted using enroßoxacin (Sigma, cat. no. 17849) at a Þnal concentration of 25 μg/ml.

Anticipated Results Using this detailed guideline and demonstration, the procedures can be easily followed by a researcher without prior VM DA isolation experience. In roller tube aggregation cultures (Alternate Protocol 1), macroscopically visible aggregates should form within two divisions. Those can be monitored under a

standard microscope at 4× to 10× magniÞcation by focusing on the aggregate that has sunk to the bottom of the conical tube. Using the adherent culture system (Alternate Protocol 2) for expansion of VM precursor cells (rat E11 embryos; CRL ∼6 mm), a net expansion of 3- to 5-fold over the course of 3 days can be expected. bFGF is used as an efÞcient mitogen for VM DA precursors (Bouvier and Mytilineou, 1995; Studer et al., 1998). On average, 60% of cells at this stage are Nestin+ when cultured with bFGF-containing medium as described here (ranging from up to 85% of cells at day 2 in culture to ∼40% after 7 days of expansion). Approximately 45% of cells incorporate the proliferation marker bromodeoxyuridine (BrdU) during the expansion phase, when incubated under the conditions described above. For VM expansion cultures, E11 embryos (CRL ∼6 mm) yield the highest efÞciency/number of cells. Note that longer expansion times result in a decrease in proliferative capacity (e.g., 30% BrdU+ cells at 7 div). Expect ∼90% of cells to be positive for markers of neuronal phenotypes at this stage. Contaminating astrocytes, positive for glial markers such as GFAP, are routinely 1 month after preparation.

Anticipated Results This protocol is used to establish karyotypically normal NSC lines that are stable regarding their growth kinetics, self renewal, and differentiation potential. For adult mouse SVZ NSCs, each subculturing step results in a twoto five-fold increase in cell number (up to tenfold for mouse embryonic NSC and two-fold for human fetal NSCs). As the initial number of NSCs in the dissociated tissue may depend on many variables as described above, a minimum of 106 cells (for mouse tissue) and 5 × 105 cells (for human tissue) is expected 2 weeks after the initial tissue dissociation.



Time Considerations Basic Protocol 1 It takes 40 min to isolate SVZ regions from three animals or one region from embryos deriving from each pregnant mouse. Enzymatic digestion of tissue takes 30 to 40 min. Its requires 30 min to set up an NSC primary culture. Basic Protocol 2 It takes 15 min to subculture each flask. To establish a stable NSC line from adult mouse brain, 1 month of passaging is required, 20 days from embryonic mouse brain, 2 months from human fetal brain. Basic Protocol 3 For adhesive substrate preparation, 2 hr are required and 30 min are required to prepare dissociated neurospheres and plate a complete 24/48-well plate. Basic Protocol 4 To prepare a complete 96-well plate, 30 min are required. Alternate Protocol 1 It takes 20 min to prepare methyl-cellulose gel matrix, 30 min for cell harvesting and counting, and 10 min for seeding a petri dish. Alternate Protocol 2 Thirty minutes are required for subcloning at each passage. Basic Protocol 5 It takes 15 min to collect neurospheres into the cryovial. It takes an additional ∼6 hr to move the cryovial to liquid nitrogen.

Literature Cited Bjornson, C.R., Rietze, R.L., Reynolds, B.A., Magli, M.C., and Vescovi, A.L. 1999. Turning brain into blood: A hematopoietic fate adopted by adult neural stem cells in vivo. Science 283:534-537. Bottai, D., Fiocco, R., Gelain, F., Defilippis, L., Galli, R., Gritti, A., and Vescovi, L.A. 2003. Neural stem cells in the adult nervous system. J. Hematother. Stem Cell Res. 12:655-670. Cedrola, S., Guzzi, G., Ferrari, D., Gritti, A., Vescovi, A.L., Pendergrass, J.C., and La Porta, C.A. 2003. Inorganic mercury changes the fate of murine CNS stem cells. FASEB J. 17:869871. Craig, C.G., Tropepe, V., Morshead, C.M., Reynolds, B.A., Weiss, S., and van der Kooy, D. 1996. In vivo growth factor expansion of endogenous subependymal neural precursor cell

populations in the adult mouse brain. J. Neurosci. 16:2649-2658. Davis, A.A. and Temple, S. 1994. A self-renewing multipotential stem cell in embryonic rat cerebral cortex. Nature 372:263-266. De Filippis, L., Lamorte, G., Snyder, E.Y., Malgaroli, A., and Vescovi, A.L. 2007. A novel, immortal, and multipotent human neural stem cell line generating functional neurons and oligodendrocytes. Stem Cells 25:2312-2321. De Filippis, L., Ferrari, D., Rota Nodari, L., Amati, B., Snyder, E., and Vescovi, A.L. 2008. Immortalization of human neural stem cells with the c-myc mutant T58A. PLoS ONE 3:e3310. Flax, J.D., Aurora, S., Yang, C., Simonin, C., Wills, A.M., Billinghurst, L.L., Jendoubi, M., Sidman, R.L., Wolfe, J.H., Kim, S.U., and Snyder, E.Y. 1998. Engraftable human neural stem cells respond to developmental cues, replace neurons, and express foreign genes. Nat. Biotechnol. 16:1033-1039. Galli, R., Borello, U., Gritti, A., Minasi, M.G., Bjornson, C., Coletta, M., Mora, M., De Angelis, M.G., Fiocco, R., Cossu, G., and Vescovi, A.L. 2000. Skeletal myogenic potential of human and mouse neural stem cells. Nat. Neurosci. 3:986991. Galli, R., Fiocco, R., De Filippis, L., Muzio, L., Gritti, A., Mercurio, S., Broccoli, V., Pellegrini, M., Mallamaci, A., and Vescovi, A.L. 2002. Emx2 regulates the proliferation of stem cells of the adult mammalian central nervous system. Development 129:1633-1644. Gritti, A., Cova, L., Parati, E.A., Galli, R., and Vescovi, A.L. 1995. Basic fibroblast growth factor supports the proliferation of epidermal growth factor-generated neuronal precursor cells of the adult mouse CNS. Neurosci. Lett. 185:151-154. Gritti, A., Parati, E.A., Cova, L., Frolichsthal, P., Galli, R., Wanke, E., Faravelli, L., Morassutti, D.J., Roisen, F., Nickel, D.D., and Vescovi, A.L. 1996. Multipotential stem cells from the adult mouse brain proliferate and self-renew in response to basic fibroblast growth factor. J. Neurosci. 16:1091-1100. Gritti, A., Frolichsthal-Schoeller, P., Galli, R., Parati, E.A., Cova, L., Pagano, S.F., Bjornson, C.R., and Vescovi, A.L. 1999. Epidermal and fibroblast growth factors behave as mitogenic regulators for a single multipotent stem celllike population from the subventricular region of the adult mouse forebrain. J. Neurosci. 19:32873297. Gritti, A., Bonfanti, L., Doetsch, F., Caille, I., Alvarez-Buylla, A., Lim, D.A., Galli, R., Verdugo, J.M., Herrera, D.G., and Vescovi, A.L. 2002. Multipotent neural stem cells reside into the rostral extension and olfactory bulb of adult rodents. J. Neurosci. 22:437445. Kilpatrick, T.J. and Bartlett, P.F. 1995. Cloned multipotential precursors from the mouse cerebrum require FGF-2, whereas glial restricted

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precursors are stimulated with either FGF-2 or EGF. J. Neurosci. 15:3653-3661.

adult human brain contains neural stem cells but lacks chain migration. Nature 427:740-744.

Louis, S.A., Rietze, R.L., Deleyrolle, L., Wagey, R.E., Thomas, T.E., Eaves, A.C., and Reynolds, B.A. 2008. Enumeration of neural stem and progenitor cells in the neural colony-forming cell assay. Stem Cells 26:988-996.

Seri, B., Herrera, D.G., Gritti, A., Ferron, S., Collado, L., Vescovi, A., Garcia-Verdugo, J.M., and Alvarez-Buylla, A. 2006. Composition and organization of the SCZ: A large germinal layer containing neural stem cells in the adult mammalian brain. Cereb. Cortex 16:i103-i111.

Qian, X., Davis, A.A., Goderie, S.K., and Temple, S. 1997. FGF2 concentration regulates the generation of neurons and glia from multipotent cortical stem cells. Neuron 18:81-93. Reynolds, B.A. and Weiss, S. 1992. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707-1710. Reynolds, B.A. and Weiss, S. 1996. Clonal and population analyses demonstrate that an EGFresponsive mammalian embryonic CNS precursor is a stem cell. Dev. Biol. 175:1-13. Reynolds, B.A. and Rietze, R.L. 2005. Neural stem cells and neurospheres—Re-evaluating the relationship. Nat. Methods 2:333-336. Ryder, E.F., Snyder, E.Y., and Cepko, C.L. 1990. Establishment and characterization of multipotent neural cell lines using retrovirus vector-mediated oncogene transfer. J. Neurobiol. 21:356-375. Sah, D.W., Ray, J., and Gage, F.H. 1997. Bipotent progenitor cell lines from the human CNS. Nat. Biotechnol. 15:574-580. Sanai, N., Tramontin, A.D., Quinones-Hinojosa, A., Barbaro, N.M., Gupta, N., Kunwar, S., Lawton, M.T., McDermott, M.W., Parsa, A.T., ManuelGarcia Verdugo, J., Berger, M.S., and AlvarezBuylla, A. 2004. Unique astrocyte ribbon in

Vescovi, A.L., Gritti, A., Galli, R., and Parati, E.A. 1999a. Isolation and intracerebral grafting of nontransformed multipotential embryonic human CNS stem cells. J. Neurotrauma 16:689693. Vescovi, A.L., Parati, E.A., Gritti, A., Poulin, P., Ferrario, M., Wanke, E., Frolichsthal-Schoeller, P., Cova, L., Arcellana-Panlilio, M., Colombo, A., and Galli, R. 1999b. Isolation and cloning of multipotential stem cells from the embryonic human CNS and establishment of transplantable human neural stem cell lines by epigenetic stimulation. Exp. Neurol. 156:71-83. Villa, A., Snyder, E.Y., Vescovi, A., and MartinezSerrano, A. 2000. Establishment and properties of a growth factor-dependent, perpetual neural stem cell line from the human CNS. Exp. Neurol. 161:67-84. Weiss, S., Dunne, C., Hewson, J., Wohl, C., Wheatley, M., Peterson, A.C., and Reynolds, B.A. 1996a. Multipotent CNS stem cells are present in the adult mammalian spinal cord and ventricular neuroaxis. J. Neurosci. 16:75997609. Weiss, S., Reynolds, B.A., Vescovi, A.L., Morshead, C., Craig, C.G., and van der Kooy, D. 1996b. Is there a neural stem cell in the mammalian forebrain? Trends Neurosci. 19:387-393.

Isolation of Neural Stem Cells from Neural Tissues



Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

UNIT 2E.1

Yuzo Niki1 1

Ibaraki University, Ibaraki, Japan

ABSTRACT This unit describes how to collect, culture, and establish stable cell lines of ovarian somatic and germline stem cells of Drosophila. We also describe a protocol for culturing embryonic cells that overexpress growth factors, which serve as a source for conditioned C 2009 by John Wiley & Sons, medium. Curr. Protoc. Stem Cell Biol. 10:2E.1.1-2E.1.9. Inc. Keywords: Drosophila r somatic and germline stem cells r isolation r expansion

INTRODUCTION Germline stem cells (GSCs) and their niches have been extensively studied in vivo in Drosophila. The concept that stem cells are controlled by particular microenvironments known as niches has been widely suggested by various in vivo approaches (reviewed by Fuller and Spradling, 2007). In vitro systems, however, are powerful and indispensable for analyzing the interactions between GSCs and their niches directly and biochemically (reviewed by Niki, 2008). Thus, this unit describes how to collect, culture, and establish stable cell lines of ovarian somatic stem (OSS) cells and GSCs from tumorous mutant ovaries. NOTE: The following tissue culture procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All incubations are performed at 25◦ C and do not need any special equipment such as a CO2 incubator. NOTE: Most fly laboratories add live (dry) yeast as food to raise flies. It is hard to remove or kill yeast by treating with any kind of antibiotics and antimicrobial drugs for tissue and cell cultures. Instead of live yeast, we feed “microwaved” yeast (see recipe). NOTE: Storing many ovaries at once in a depression glass slide frequently results in contamination with microorganisms. To reduce the risk of contamination, it is recommended that several ovaries from only one or two females be collected in a depression glass slide.

ISOLATION AND CULTURE OF OVARIAN SOMATIC STEM CELLS AND GERMLINE STEM CELLS

BASIC PROTOCOL

The protocol below describes a common procedure for collecting and culturing OSS cells and GSCs. To obtain sufficient numbers of OSS cells and GSCs, we used females of either the w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ] or w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ) P[vasa-egfp] genotype. These constructs have the wild-type allele of a bam gene ligated with a heat shock promoter. Germline cells are marked with ovo-lacZ and vasa-egfp. In these bam homozygous females, OSS cells and GSCs expand as the female ages. Somatic Stem Cells Current Protocols in Stem Cell Biology 2E.1.1-2E.1.9 Published online September 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc02e01s10 C 2009 John Wiley & Sons, Inc. Copyright


Materials 20- to 30-day-old female flies of w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ] or w1118 ; P[w+ hsp-70 bam+ ] 11-d bam86 ry e/bam86 P[ovo-lacZ) P[vasa-egfp] genotype (Fig. 2E.1.1A,B) 70% (v/v) ethanol Drosophila phosphate-buffered saline (PBS; Robb, 1969), sterile Culture medium (see recipe) Penicillin/streptomycin (see recipes), optional Distilled water DMSO Liquid nitrogen 15-ml conical tubes Single concave depression glass slides Forceps Sterilized tungsten needles 96-well tissue culture plates Sealed container (e.g., Tupperware) Phase-contrast and fluorescent microscope 200-μl pipet tips Cryotubes (Nunc) −20◦ and −80◦ C freezers 1.5-ml microcentrifuge tubes Isolate female GSCs (fGSCs) from bag-of-marbles (bam) ovaries 1. Sterilize 20- to 30-day-old female flies homozygous for the bam mutation, as described above, with 5 ml 70% ethanol for 10 min in a 15-ml conical tube. 2. Immerse several adult flies in a drop of sterilized PBS on a single concave depression glass slide. 3. Using forceps, dissect the ovaries carefully so as to not injure the gut. 4. Wash the dissected ovaries three times with culture medium. If necessary, add a mixture of penicillin and streptomycin to the culture medium. 5. Dissociate ovaries into ovarioles and then fragment each ovariole into pieces with fine tungsten needles. 6. Remove any membranous debris carefully. 7. Wash the cell masses three times, each time with 100 μl culture medium.

Plate the cells 8. Dispense 100 μl of the culture medium–containing cell masses into interior wells of a 96-well tissue culture plate. 9. Add new culture medium to make a final volume of 200 μl in each well. 10. Add 200 ml of distilled water in the external marginal wells of a 96-well culture plate to reduce the evaporation of the culture medium. 11. Store the culture plates in a sealed container, such as a Tupperware container, at 25◦ C. Culturing Ovarian Somatic and Germline Stem Cells of Drosophila

12. Check the cells with a phase-contrast microscope daily and exchange half of the culture medium every week.



A

B

C

D

E

Figure 2E.1.1 (A) A pair of tumorous bam ovaries from a 30-day-old female Drosophila melanogaster flies. (B) A living bam ovariole showing vasa-gfp positive GSCs. (C) A stable line named the fGS/OSS line. Female GSCs are located on the somatic cells that originated from follicle stem cells. (D) Cellular clump formed after confluence. (E) Living OSS cells stained with Hoechst 33248. (F) OSS cells in the subconfluent condition. Bar = 50 μm.

13. Scrape cellular clumps with a 200-μl pipet tip and disperse them by gentle pipetting. Split the cells 1:2 or 1:4 into new wells of the same culture plate when they become confluent. For expanding cells, it is much better to transfer cells into new wells than to leave them in old wells. Once split, cells continue to divide and expand rapidly thereafter. It takes ∼20 hr for the doubling of cell number. Split cultured cells before they become confluent. Note that GSCs are sensitive to cell density and sometimes disappear rapidly if you leave the cells alone after they have become confluent (Fig. 2E.1.1D). During primary culture, the cells start to spread and continue to divide for 1 to 2 weeks after initiation of culture, but cease to divide thereafter. It is important to continue to exchange the medium once every week even if the cultured cells have ceased to grow and turned brown or black, a symptom of cell death. There will be cases in which several transparent cells appear at the periphery of the colored cellular aggregates and begin to divide again one or several months after culture. These somatic cells originate from

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daughter cells of the somatic stem cells. The cells then start expanding dramatically and cover the bottom of the culture well to form a sheet. There will also be cases in which round, large cells, characteristic of GSCs, appear in the somatic sheet. GSCs associated with somatic cells continue to divide as the somatic sheet expands (Fig. 2E.1.1C).

Establish cell lines consisting of only ovarian somatic cells 14. Seed 1 to 50 cells, from the above GSC/OSS cell culture, in each well of a 96-well plate separately, after dissociating with a 200-μl tip. 15. Select wells consisting only of somatic cells by checking the vasa-EGFP activity in the live condition with a phase-contrast and fluorescent microscope. 16. Repeat the dilution and expansion of the somatic cells several times until the somatic cells show the same phenotype in all subpopulations (Fig. 2E.1.1E,F).

Freeze and thaw cultured cells 17. Add 20 μl DMSO to the well and gently disperse the cells by pipetting. Transfer 1–10 × 104 cells into a cryotube. 18. Store the cells overnight at −20◦ C. 19. Transfer the cells to a −80◦ C deep freezer and then store them in liquid nitrogen. 20. Transfer the cryotube to a clean bench and warm the tubes with water at 25◦ C to thaw the frozen cells. 21. Wash the cells with 400 μl culture medium, transfer them into a 1.5-ml microcentrifuge tube, and remove the DMSO by centrifuging 1 min at 500 × g in a microcentrifuge at 25◦ C. 22. Seed 1–10 × 103 cells into the wells of a culture plate. 23. Add fresh culture medium to make the final volume 200 μl in each well.

Phenotype the cells 24. Discriminate GSCs and OSS cells from each other by size and morphology: GSCs are round and 10-μm in diameter and are usually located on the OSS cell sheet (Fig. 2E.1.1C), whereas OSS cells are small and flattened and are 50 ml per whole intestine): 25% ethanol, 15 min at room temperature 50% ethanol, 15 min at room temperature 75% ethanol, 15 min (or overnight at 4◦ C for large pieces) 90% ethanol, 30 min at room temperature 100% ethanol, three times, 1 hr each at room temperature. 6. Replace the last ethanol wash with xylene (>50 ml) and incubate 1 hr at room temperature. Repeat xylene treatments twice, for a total of three times. 7. Place the samples in parafÞn wax (>100 ml). Replace once and incubate overnight in a 55◦ C oven. 8. Replace the parafÞn once and allow samples to solidify at room temperature. 9. Prepare 8- to 10-μM sections and mount on adhesive glass slides. 10. Dry the slides overnight in a 45◦ C oven.

Pretreat sections prior to in situ hybridization 11. Clean glass jars (including covers) suitable for holding glass slides (e.g., Coplin jars with lids) and bake overnight at 200◦ C. 12. Proceed with standard dewaxing and rehydration protocol, by placing slides in jars containing the following solutions for the indicated times:

Xylene, three times, 5 min each 100% ethanol, two times, 5 min each 96% ethanol, one time, 5 min 70% ethanol, one time, 5 min 50% ethanol, one time, 5 min 25% ethanol, one time, 5 min. For each step listed above, it is important to add enough solution in the jar to cover intestinal sections. The exact volumes will depend on the size and shape of jars and the number of slides. Do not let sections dry out during any step.

13. Rinse the slides twice in DEPC-treated H2 O. 14. Treat the slides with 0.2 N HCl for 15 min at room temperature.

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Table 2F.1.1 Incubation Times, Concentration, and Temperature of Proteinase K Treatment for Various Types of Intestinal Sections

Tissue type

Proteinase K concentration in PBS

Time

Temperature

Embryo, E8.5

15 μg/ml

5 min

Room temperature

Embryo, E10.5

15 μg/ml

10 min

Room temperature

Fetal gut, E14.5-E18.5

30 μg/ml

10 min

Room temperature

Adult intestine and colon

30 μg/ml

20 min

37◦ C

15. Incubate the sections with proteinase K in PBS buffer (see Table 2F.1.1 for concentration, temperature, and durations). Place jar with slides in a water bath if required. Optimal concentration and duration of proteinase K treatment should be tested carefully.

16. Rinse the slides in freshly prepared 0.2% glycine/PBS solution for 1 min. Glycine should be added to PBS solution at last moment. Do not store solution for long periods of time.

17. Rinse the slides twice, each time for 1 min in PBS. 18. Post-Þx for 10 min with 4% paraformaldehyde in PBS at room temperature. 19. Rinse the slides three times, each time for 1 min in PBS. 20. Prepare fresh acetic anhydride solution. Shake vigorously and add to the slides in a glass jar. Incubate for 5 min at room temperature. 21. Repeat acetic anyhydride treatment once. 22. Rinse Þve times, each time in PBS for 1 min at room temperature. 23. Rinse two times in 5× SSC, pH 7.5, each time for 1 min at room temperature.

Prehybridize the sections 24. Remove excess solution from slides with tissue and place them in a covered slide box humidiÞed with 5× SSC, pH 7.5/50% formamide. 25. Add enough hybridization solution (∼500 μl) to completely cover the sections. It is not necessary to place a coverslip over section.

26. Incubate the slide box in a 65◦ C oven for at least 1 hr.

Hybridize the sections 27. Remove excess hybridization solution and replace with 300 to 400 μl/slide of hybridization solution containing 500 ng/ml of digoxigenin-labeled probe. Take care that the slides are horizontal in the humidiÞed chamber. Again, no coverslips are necessary.

28. Incubate the slides in an oven at 62◦ to 70◦ C for 24 to 72 hr. Optimal incubation times and temperatures should be empirically tested, although 48 hr at 65◦ C gives satisfactory results for most probes.

In Situ Hybridization to Identify Gut Stem Cells

Wash post-hybridization 29. Remove excess hybridization solution and place in glass jar. Rinse in 2× SSC, pH 7.5, for 1 min at room temperature.



30. Wash the slides three times for 20 min each at 60◦ to 65◦ C in 50% formamide/2× SSC, pH 4.5. Optimal washing temperature should be tested.

31. Rinse the slides Þve times, each time in Tris/NaCl buffer for 1 min at room temperature.

Detect signal immunologically 32. Remove excess solution from the slides with tissue and place them in a covered slide box humidiÞed with water. 33. Apply blocking solution over sections (500 μl per slide) and incubate at room temperature for at least 30 min. 34. Dilute sheep anti-digoxigenin antibody 1/2000 in blocking solution. 35. Remove blocking solution and replace with antibody solution (400 μl per slide). Incubate overnight or longer at 4◦ C. 36. Wash the slides Þve to seven times, each time in Tris/NaCl buffer for 1 min at room temperature. 37. Wash the slides two to three times, each time in NTM buffer for 1 min at room temperature. 38. Add NBT/BCIP working solution to sections in a humidiÞed slide box. Incubate for up to 24 hr at room temperature, keeping the slides in the dark. 39. Wash the slides twice, each time in PBS for 1 min at room temperature. 40. Dehydrate the sections as follows:

Rinse sections in H2 O Rinse in 70% (v/v) ethanol Rinse in 90% (v/v) ethanol Rinse twice in 100% (v/v) ethanol Rinse twice in xylenes. Proceed quickly through each step. Part of the signal may be lost with extensive washes in ethanol. To avoid signal loss, sections may be visualized under the microscope prior to dehydration. In this case, a few drops of glycerol are added to the slide followed by a coverslip. This method will also reduce the diffusion of the signal resulting from dehydration in ethanol.

41. Apply mounting medium and place a coverslip over the section. 42. Examine the slides with a light microscope. See Anticipated Results for a description of a positive signal.

GENERATION OF DIGOXIGENIN RNA PROBES The generation of digoxigenin RNA probes is achieved by an in vitro transcription reaction of linearized template DNA using T7, T3, or SP6 RNA polymerases. During the in vitro transcription reaction, digoxigenin-coupled UTPs are incorporated into the RNA probe. To generate anti-sense probes that will recognize sense mRNA, template DNA is cut using a restriction enzyme that creates a 5 overhang (avoid 3 overhangs) at the 5 end of the cDNA. Ensure that following the digestion the T7, T3, or SP6 promoter is at the 3 end of the template DNA. Accordingly, to generate sense probes, template DNA can be cut at the 3 end.

SUPPORT PROTOCOL

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A useful source of probes for in situ hybridization can be found through the IMAGE consortium (http://image.hudsonalpha.org/). Using Web-based tools speciÞc cDNAs can be searched and ordered. For detecting stem cells, we recommend generating a full-length probe of Olfm4. When compared to other stem cell markers (e.g., Lgr5, Ascl4, or Bmi1), Olfm4 gives strong and highly reproducible signals, and for this reason, it is perhaps the most useful marker to detect Lgr5+ stem cells by ISH.

Materials Plasmid for gene of interest (e.g., see Table 2F.1.3) Restriction endonuclease and buffer (Bloch and Grossman, 1995) Agarose 3 M sodium acetate, pH 5.2 Phenol/choloroform Absolute ethanol 70% (v/v) ethanol 10 × transcription buffer (Roche) Dithiothreitol (DTT) 10 × Dig RNA labeling mix (Roche) RNase inhibitor (Fermentas) T7 or T3 or SP6 RNA polymerases (Roche) DEPC-treated H2 O Rnase-free DNaseI (Fermentas), optional 4 M LiCl Formamide 1.5-ml microcentrifuge tubes RNA puriÞcation columns (RNeasy Mini Kit, Qiagen) Additional reagents and equipment for agarose gel electrophoresis (Voytas, 2000) and digestion of DNA with restriction enzymes (Bloch and Grossman, 1995) Digest plasmid DNA 1. Digest 10 μg of plasmid DNA with the appropriate restriction enzyme (Bloch and Grossmann, 1995). 2. Ensure that template DNA is completely linearized by running an aliquot on 1% agarose gel (Voytas, 2000). 3. Add 1/10 volume of 3 M sodium acetate, pH 5.2, to the digest and extract once with 50:50 (v/v) phenol/chloroform mixture and once again with chloroform to remove trace amounts of phenol. 4. Precipitate DNA by adding 2.5 vol of absolute ethanol and centrifuge 10 min at 14,000 rpm, 4◦ C, in a benchtop microcentrifuge. 5. Wash the pellet in 500 μl of 70% ethanol and resuspend in 15 μl water. To avoid phenol/chloroform extraction and ethanol precipitation steps, linearized DNA can be puriÞed using commercially available columns.

Perform in vitro translation with digoxigenin dUTP 6. Prepare in vitro transcription reaction in a 1.5-ml microcentrifuge tube (20-μl reaction volume) as follows: In Situ Hybridization to Identify Gut Stem Cells

1 to 2 μg of linearized DNA (from step 5) 1× transcription buffer (usually supplied by manufacturer of RNA polymerases) 2 μl of 0.1 M DTT 2 μl 10× Dig RNA labeling mix



20 U of RNase inhibitor 30 U of T7 or T3 or SP6 RNA polymerase. Use the polymerase that corresponds to the promoter at the 3 end of the linearized DNA template. See the Introduction above.

7. Incubate the reaction at 37◦ C for 3 hr. 8. Clean-up cRNA products by using commercially available RNA puriÞcation columns. Elute samples from columns with 50 μl DEPC-treated H2 O. As an alternative to using RNA puriÞcation columns researchers may follow these steps to clean up cRNA reaction. a. Add 1 U RNase-free DNaseI to the in vitro transcription reaction, and incubate samples 15 min at 37◦ C. b. Add 2.5μl 4 M LiCl and 75 μl ethanol to the reaction; store at −70◦ C for 20 min. c. Microcentrifuge 5 min at 14,000 rpm, 4◦ C. Wash the pellet in 500 μl of 70% ethanol and resuspend in 50 μl DEPC-treated H2 O.

9. Remove 1 μl of the puriÞed probe to measure the concentration and 3 μl for electrophoresis on standard 1% agarose/ethidium bromide gel (Voytas, 2000). Add an equal volume of 100% formamide to the remaining probe and store at −70◦ C.


Acetic anhydride solution 0.25% (v/v) acetic anhydride in 0.1 M triethanolamine, pH 8.0 Add acetic anhydride solution immediately before use. Do not store! 0.1 M triethanolamine, pH 8.0, may be stored at room temperature

Blocking solution 1% (w/v) blocking powder (Roche) in 1× Tris/NaCl buffer (see recipe). Heat at 65◦ C to dissolve. Usually prepared fresh, but can be stored up to 3 months at −20◦ C.

Hybridization solution 50% (v/v) formamide 5× SSC, pH 4.5 2% (w/v) blocking powder (Roche) 0.05% (w/v) CHAPS 5 mM EDTA 50 μg/ml heparin 1 μg/ml yeast RNA Heat at 65◦ C to dissolve Usually prepared fresh, but can be stored up to 3 months at −20◦ C.

NBT/BCIP solution NBT stock solution: 10 mg of NBT (Sigma) in 1 ml of H2 O continued

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Heat and vortex if required Protect from light and store up to 6 months at −20◦ C BCIP stock solution: 25 mg of BCIP (Sigma) in 500 μl DMF (dimethylformamide) Store up to 6 months at −20◦ C Working solution: 10 ml NTM buffer (see recipe) 25 μl 1 M levamisole 333 μl NBT stock solution 35 μl BCIP stock solution Prepare fresh NTM buffer 0.1 M Tris·Cl, pH 9.5 0.1 M NaCl 0.05 M MgCl2 Prepare fresh Paraformaldehyde (PFA) in PBS, 4% (w/v) Heat at 65◦ C to dissolve Prepare fresh Phosphate-buffered saline (PBS) For 1 liter: 8 g NaCl 0.2 g KCl 2.68 g Na2 HPO4 ·7H2 O 0.24 g KH2 PO4 800 ml H2 O Adjust pH to 7.4 with HCl Adjust volume to 1 liter with H2 O Store up to 6 months at room temperature SSC, 20× For 1 liter: 175.3 g NaCl 88.2 g sodium citrate·2H2 O 800 ml H2 O Adjust pH to 7.5 or 4.5 with HCl Adjust volume to 1 liter with H2 O Store up to 6 months at room temperature When SSC is combined with formamide, pH 4.5 is used. This will ensure that the formamide/ SSC solution will be pH neutral.

Tris/NaCl buffer

In Situ Hybridization to Identify Gut Stem Cells

0.1 M Tris·Cl, pH 7.5 0.15 M NaCl 0.1 % (v/v) Tween 20 Store up to 6 months at room temperature



COMMENTARY Background Information More than 30 years ago, Cheng and Leblond proposed that intestinal stem cells are located at the base of the crypts of Lieberkühn (Cheng and Leblonde, 1974a,b). These socalled crypt base columnar (CBC) cells were found to be highly proliferative cells, capable of giving rise to the four differentiated cell types of the intestinal epithelium (i.e., enterocytes, goblet cells, Paneth cells, and enteroendocrine). Around the same time, however, pulse-chase experiments with tritiated thymidine led Potten and colleagues to different conclusions. Indeed these researchers demonstrated that label-retaining cells were preferentially located immediately above the Paneth cell compartment (Potten et al., 1974, 2002; Potten, 1977). This population of stem cells is referred to as +4 cells given their position along the crypt-villus axis relative to the bottom of the crypts. Although several question marks still persist regarding the properties of these two populations (+4 cells versus CBC cells), recent knock-in strategies have led investigators in the Þeld to more reliably identify and track intestinal stem cells. In the Þrst model, Barker et al. labeled CBC cells using mice engineered to carry a GFP and a tamoxifen inducible Cre in the Lgr5 locus (Barker et al., 2007). In follow up studies, the Lgr5-GFP-CreER allele was utilized to identify novel stem cell markers based on transcriptional proÞling of sorted GFP-expressing CBC cells (van der Flier et al., 2009). A similar approach allowed Sangiorgi and Capechi to speciÞcally label +4 cells by introducing the CreER enzyme in the locus of the Polycomb family member, Bmi1 (Sangiorgi and Capecchi, 2008). In either case, crossing these lines with the R26RLacZ reporter mice to permanently mark +4 cells or CBC cells demonstrated unequivocally that both cell populations are long-lived and generate all four cell types of intestinal epithelium.

Critical Parameters and Troubleshooting The optimal length of the probe should be determined empirically. In most cases, fulllength probes give satisfactory results. Note however that the yield and quality of the in vitro transcription reaction may be limited when using very large template DNA (>3 kB). PCR fragments with the appropriate promoter added to the 3 end can also be used as an alter-

native to plasmid DNA for generating template DNA. Although alkaline phosphatase (AP)–based detection schemes offer high sensitivity, one caveat in the intestine is the fact that the epithelium harbors a high level of endogenous AP activity. To circumvent this problem it is important to inactivate endogenous AP activity by pretreating sections in 0.2 N HCl. The drug levamisole is an effective way to reduce endogenous AP activity when performing in situ hybridization on whole embryos. In the case of adult intestinal material, however, levamisole is not sufÞcient to block AP activity. Incomplete inactivation of enodogenous AP appears as a thin apical staining of enterocytes. It is also believed that HCl treatments may favor accessibility of the probe to target sequences by extracting mRNA-bound proteins. Another commonly used treatment is acetylation, using acetic anhydride (0.25%) in triethanolamine. This treatment is also thought to be important for decreasing background, but it also appears to inactivate RNases and may help in producing a stronger signal. For troubleshooting information for in situ hybridization, see Table 2F.1.2.

Anticipated Results For probe synthesis, cRNA samples run on normal agarose gels usually give a prominent but diffused band. Occasionally, certain probes may give more of a smear. One can expect to obtain a yield of 7 to 15 μg of cRNA. Following ISH, the intensity and speciÞcity of the signal will depend on the expression pattern and levels of the gene of interest. Generally, for the intestine, genes that are expressed in differentiated epithelial cell types (enterocytes, goblet cells, enteroendocrine cells, and Paneth cells) are readily detected within a few hours of development. Genes exclusively expressed in progenitor/stem cells may require longer development times and signal may be weaker. Generally, the staining pattern appears as a uniform blue or purple deposit within the contours of a given cell. Stem cell markers should result in discrete staining of CBC cells and/or +4 cells and no staining should be observed in the adjacent Paneth cells. The latter can be easily distinguished from CBC or +4 cells by their large size and granular morphology. Occasionally, staining will appear as discrete dots in or around the Paneth cells. This type of pattern should be considered as background.

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Table 2F.1.2 Troubleshooting Guide to In Situ Hybridization to Identify Intestinal Stem Cells

Problem

Possible cause

Solution

Poor histology

Fixation inadequate

Make sure intestines are immediately Þxed following dissection. Take care when ßushing intestines or ßush directly with formalin instead of PBS. Reduce time and concentration of proteinase K treatment

Proteinase K treatment too long Very strong and spotty staining in Paneth, goblet, or enteroedocrine cells

UnspeciÞc binding to secretory products of Paneth, goblet, or enteroendocrine cells

Prepare new intestinal sections. Use a different probe.

Weak or no staining

Temperature of hybridization or post-hybridization washes is too stringent

Prepare new intestinal sections. Lower hybridization/washing temperature. Use a different probe.

High background

Temperature of hybridization or post-hybridization washes is not stringent enough

Raise hybridization/washing temperature. Use a different probe.

Table 2F.1.3 Useful Epithelial, Cell-Type SpeciÞc Markers for In Situ Hybridization in the Small Intestine

Cell type

Gene

Enterocytes

Fabp1, Fabp2

Goblet cells

Gob5, Tff3

Enteroendocrine cells

Chromogranin B

Paneth cells

Cryptdins

Entire epithelium

Villin, Tcf4

Proliferative crypt epithelial cells

c-Myc, c-Myb

Early progenitors/stem cells

Axin2, Lgr5, Olfm4

Table 2F.1.3 lists genes from which suitable probes can be generated to mark certain cell types of the intestine.

Time Considerations The generation of labeled probe including digestion and in vitro transcription reactions can be completed in a single day or a day and a half. The Þxation, dehydration, and parafÞn embedding of intestinal samples require at least 24 to 36 hr. The in situ hybridization protocol takes a minimum of 4 days depending on the length of hybridization and duration of staining procedure (see individual steps).

Literature Cited In Situ Hybridization to Identify Gut Stem Cells

Barker, N., van Es, J.H., Kuipers, J., Kujala, P., van den Born, M., Cozijnsen, M., Haegebarth, A., Korving, J., Begthel, H., Peters, P.J., and Clevers, H. 2007. IdentiÞcation of stem cells in

small intestine and colon by marker gene lgr5. Nature 449:1003-1007. Bloch, K.D. and Grossmann, B. 1995. Digestion of DNA with restriction endonucleases. Curr. Protoc. Mol. Biol. 31:3.1.1-3.1.21. Cheng, H. and Leblond, C.P. 1974a. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. I. Columnar cell. Am. J. Anat. 141:461-479. Cheng, H. and Leblond, C.P. 1974b. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. V. Unitarian theory of the origin of the four epithelial cell types. Am. J. Anat. 141:537561. Potten, C.S. 1977. Extreme sensitivity of some intestinal crypt cells to x and gamma irradiation. Nature 269:518-521. Potten, C.S., Kovacs, L., and Hamilton, E. 1974. Continuous labelling studies on mouse skin and intestine. Cell Tissue Kinet. 7:271-283.



Potten, C.S., Owen, G., and Booth, D. 2002. Intestinal stem cells protect their genome by selective segregation of template DNA strands. J. Cell Sci. 115:2381-2388. Sangiorgi, E. and Capecchi, M.R. 2008. Bmi1 is expressed in vivo in intestinal stem cells. Nat. Genet. 40:915-920. van der Flier, L.G., van Gijn, M.E., Hatzis, P., Kujala, P., Haegebarth, A., Stange, D.E., Begthel, H., van den Born, M., Guryev, V., Oving, I., van Es, J.H., Barker, N., Peters, P.J., van der Wetering, M, and Clevers, H. 2009. Transcription factor achaete scute-like 2 controls intestinal stem cell fate. Cell 136:903-912. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9.

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Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

UNIT 2G.1

Ivan Bertoncello1 and Jonathan McQualter1 1

Lung Regeneration Laboratory, The Department of Pharmacology, University of Melbourne, Victoria, Australia

ABSTRACT Adult mouse lung epithelial stem/progenitor cells (EpiSPC) can be defined in vitro as epithelial colony-forming units that are capable of self-renewal, and which when cocultured with lung mesenchymal stromal cells (MSC) are able to give rise to differentiated progeny comprising mature lung epithelial cells. This unit describes a protocol for the prospective isolation and in vitro propagation and differentiation of adult mouse lung EpiSPC. The strategy used for selection of EpiSPC and MSC from adult mouse lung by enzymatic digestion and flow cytometry is based on the differential expression of CD45, CD31, Sca-1, EpCAM, and CD24. The culture conditions required for the differentiation (co-culture with MSC) and expansion (stromal-free culture with FGF-10 and HGF) of C 2011 by John EpiSPC are described. Curr. Protoc. Stem Cell Biol. 16:2G.1.1-2G.1.12. Wiley & Sons, Inc. Keywords: lung epithelium r stem cells r colony-forming assay

INTRODUCTION This unit describes isolation and culture of adult mouse lung epithelial stem/progenitor cells (EpiSPC). EpiSPC are defined as colony-forming units (CFU) that have the capacity for self-renewal and are able to generate progeny of differentiated lung epithelial cells. Basic Protocol 1 describes a cell-fractionation method using flow cytometry for isolating EpiSPC and mesenchymal stromal cells (MSC) from enzymatically digested adult mouse lung tissue, based on their differential expression of CD45, CD31, Sca-1, EpCAM, and CD24 (McQualter et al., 2010). Basic Protocols 2 and 3 describe culture methods for the differentiation, maintenance, and propagation of adult lung EpiSPC, respectively. When co-cultured with MSC in an organotypic three-dimensional Matrigel culture, EpiSPC generate colonies comprising mature differentiated lung epithelial cells. In stromal-free cultures, EpiSPC undergo clonal proliferation when supplemented with FGF-10 and hepatocyte growth factor (HGF), and can be enzymatically dissociated and passaged using this culture setup, demonstrating their high proliferative capacity and ability to self-renew.

ISOLATION OF EPITHELIAL STEM/PROGENITOR CELLS AND MESENCHYMAL STROMAL CELLS FROM ADULT MOUSE LUNG

BASIC PROTOCOL 1

This protocol is used for the dissociation of adult mouse lung tissue and the enrichment of cell fractions containing EpiSPC and MSC. A combination of mechanical and enzymatic dissociation is used to prepare a single-cell suspension of lung cells, followed by a pre-enrichment step using discontinuous density gradient centrifugation to remove high-density cells such as erythrocytes, mature myeloid cells, and cellular debris. Flow cytometry is then used to identify and sort subpopulations of cells enriched for either EpiSPC or MSC, based on the expression of defined cell-surface markers (McQualter et al., 2009, 2010). Somatic Stem Cells Current Protocols in Stem Cell Biology 2G.1.1-2G.1.12 Published online January 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc02g01s16 C 2011 John Wiley & Sons, Inc. Copyright

2G.1.1 Supplement 16

Materials Collagenase, type 1 (Worthington) Phosphate buffered saline (PBS; Sigma, cat. no. P-3813) Adult mice (strain C57BL/6; 6 to 12 weeks; 18 to 22 g; male or female) PBS with 2% (v/v) fetal bovine serum (FBS; JRH Biosciences, batch tested) Nycoprep density gradient medium (1.077 g/cm3 , 265 mOsm (see recipe) Fluorochrome-conjugated antibodies against: EpCAM (clone G8.8) CD24 (clone M1/69) Sca-1 (clone E13-161.7) CD45 (clone 30-F11) CD31 (clone 390) Viability dye (4 ,6-diamidino-2-phenylindole (DAPI), propidium iodide (PI), or FluoroGold (Fluorochrome LLC, http://www.fluorochrome.com) Dissecting equipment 15- and 50-ml conical polypropylene centrifuge tubes (e.g., BD Falcon) 60-mm-diameter Petri dishes Thermomixer (Eppendorf) 18-G and 21-G needles 5-ml and 20-ml syringes 40-μm cell strainer (BD Biosciences) Refrigerated centrifuge (with capacity for 5-ml, 15-ml and 50-ml tubes) Sysmex KX-21N cell counter (Sysmex Corporation; http://www.sysmex.com/) or hemacytometer counting chamber and trypan blue (UNIT 1C.3) Sterile mixing cannulas (Unomedical, cat. no. 500.11.012; http://www.unomedical.com) 5-ml FACS tubes with 35-μm cell strainer caps (BD Biosciences), sterile Flow cytometer Additional reagents and equipment for euthanasia of mice (Donovan and Brown, 2006), determining viable cell concentration (UNIT 1C.3), and flow cytometry (Robinson et al., 2010) Dissociate lung tissue 1. Prepare 1 mg/ml collagenase type I solution in sterile PBS and preheat to 37◦ C, allowing 3 ml per whole mouse lung. 2. Euthanize mouse by cervical dislocation (Donovan and Brown, 2006), open mouse abdominal cavity, and sever major arteries/vessels behind intestines to exsanguinate animal. 3. Open thoracic cavity, carefully dissect out the lungs, place into a 15-ml tube containing 10 ml chilled PBS, and agitate to rinse out excess blood. If preparing multiple lungs, pool lungs and rinse into a 50-ml tube (up to five lungs per tube). This protocol has been optimized for the isolation of distal lung cells. All upper airways are carefully removed at this point.

4. Transfer lungs into a second 60-ml tube of fresh chilled PBS and rinse lungs well by gentle agitation. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

The lungs should become pale when adequately rinsed. Rinse a third time if necessary.

5. Transfer the lungs into a sterile 60-mm-diameter Petri dish and finely mince the lungs using fine dissecting scissors or a single-sided razor blade.



Prepare single-cell suspension 6. Transfer the minced tissue into a 15-ml conical centrifuge tube and add the preheated collagenase type I solution (3 ml per lung) to the tube. If processing multiple lungs, use a 50-ml centrifuge tube (up to five lungs per tube).

7. Place the tube in a Thermomixer (Eppendorf) and agitate (750 rpm, 37◦ C) for 30 min. 8. Remove the tube from the Thermomixer and triturate with an 18-G needle attached to a 5-ml syringe until chunks of tissue are mostly dissociated. If processing multiple samples, use a 20-ml syringe.

9. Return tube to the Thermomixer and agitate (750 rpm, 37◦ C) for a further 15 to 30 min until most of the lung tissue fragments are digested. When tissue digestion is complete, you should be unable to see chunks of pink lung tissue, although clumps of extracellular matrix will remain visible as white strands in the suspension.

10. Following digestion, remove the tube from the Thermomixer and triturate with a 21-G needle attached to a 20-ml syringe to generate a single-cell suspension. 11. Strain the tissue digest through a 40-μm cell strainer into a clean, sterile 50-ml centrifuge tube. 12. Resuspend the tissue digest in 50 ml of PBS containing 2% fetal bovine serum and centrifuge 5 min at 400 × g, 4◦ C. 13. Remove supernatant and resuspend the cell pellet in 50 ml of PBS containing 2% fetal bovine serum. To maximize cell recovery, supernatant can be collected in a 50-ml tube and recentrifuged.

14. Centrifuge tube(s) 5 min at 400 × g, 4◦ C. 15. Remove supernatant(s) and resuspend the cell pellet(s) in PBS containing 2% fetal bovine serum (5 ml per lung). 16. Count the cells and calculate cell concentration. A Sysmex KX-21N automated cell counter can be used to count cells, or alternatively a hemacytometer can be used with trypan blue (UNIT 1C.3) to exclude nonviable cells. Average cell yield per lung is 2.2 × 107 cells.

Prepare low-density cell fraction 17. Transfer the cell suspension (5 ml) into a 15-ml centrifuge tube and underlay the suspension with 3 ml of Nycoprep (1.077 g/cm3 ) using a sterile mixing cannula attached to a 20-ml syringe. If processing multiple lungs, the equivalent cell suspension from four lungs (20 ml) can be transferred to a 50-ml centrifuge tube and underlayed with 10 ml of Nycoprep.

18. Centrifuge the gradients 15 min at 600 × g, 21◦ C, with the brake off. Gradient centrifugation will result in most high-density cells (i.e., erythrocytes, mature myeloid cells, and cellular debris) passing through the Nycoprep, leaving an enriched band of low-density cells at the Nycoprep interface.

19. Harvest low-density cells from the interface between the PBS layer and the Nycoprep solution using a 10-ml pipet and transfer into a 50-ml centrifuge tube (Fig. 2G.1.1A). If processing multiple lungs, the low-density cell fractions can be pooled at this point. Average low-density cell yield per lung is 6.9 × 106 cells.

Somatic Stem Cells

2G.1.3 Current Protocols in Stem Cell Biology

Supplement 16

A

B

C

104

EpCAM (647)

Viability (PI)

103 102 101 100 0

Nycoprep High-density cell pellet

EpCAM (647)

Low-density cell band

20K

FSC

E

104 103 102 101 100 100

101

102

103

104

103 102 101 100 100

30K

EpCAM (647)

D FBS-2%FBS

10K

104

101

102

103

104

CD45 + CD31 (488) 104 103 102 101 100 100

Sca-1 (PE)

101

102

103

104

CD24 (PacBlue)

Figure 2G.1.1 Cell fractionation strategy used to enrich and sort EpiSPC and MSC from enzymatically digested lung tissue. (A) Photographic image of low-density cell band from density gradient centrifugation. (B) Gating strategy inclusion of PIneg viable cells, and (C) exclusion of CD45neg CD31neg cells. (D) Gating strategy for sorting CD45neg CD31neg EpCAMneg Sca-1hi MSC. (E) Gating strategy for sorting CD45neg CD31neg EpCAMhi CD24low EpiSPC.

20. Top up tube(s) containing the low-density cells to a total volume of 50 ml with PBS containing 2% fetal bovine serum and centrifuge 5 min at 400 × g, 4◦ C. 21. Discard supernatant(s) and resuspend the cell pellet(s) in 50 ml PBS containing 2% fetal bovine serum. 22. Centrifuge the tube(s) 5 min at 400 × g, 4◦ C. 23. Discard supernatant(s) and resuspend the cell pellet(s) in 5 ml per lung of PBS containing 2% fetal bovine serum. 24. Count the cells and calculate cell concentration (UNIT 1C.3).

Prepare cells for flow cytometry 25. Aliquot 100,000 to 200,000 cells into a 5-ml FACS tube for each of the compensation tubes and 500,000 cells for isotype control tubes. Set aside remaining cells in a 15-ml conical centrifuge tube (cells for sorting). Compensation tubes should be prepared for each of the antibodies and fluorochromes to be used in the sort strategy. An aliquot of unstained cells should also be prepared.

26. Wash all samples, including cells for sorting, with PBS containing 2% fetal bovine serum, centrifuge 5 min at 400 × g, 4◦ C, and discard supernatant. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

27. Resuspend cell pellets in compensation tubes in 50 μl of each optimally pretitered antibody, and resuspend cells for sorting at 5 × 106 cells per 100 μl of optimally pretitered antibody combination. Use PBS containing 2% fetal bovine serum to dilute antibodies. If streptavidin-conjugated antibodies are used, PBS containing 0.5% (w/v) BSA should be used as a diluent to prevent reactivity of streptavidin with serum biotin.



28. Incubate all samples on ice in the dark for 20 min. 29. Wash cells in PBS containing 2% fetal bovine serum, centrifuge 5 min at 400 × g, 4◦ C, and discard supernatant. Repeat steps 26 to 29 if secondary or tertiary antibody labeling steps are required.

30. Resuspend cells in compensation tubes in 300 μl and cells for sorting and isotype control tubes at 10–15 × 106 cells/ml in PBS containing 2% fetal bovine serum and an appropriate viability dye. Depending on the flow cytometer setup and laser configuration, DAPI (0.5 μg/ml), PI (1 μg/ml), or Fluorogold (2 μM) may be used as a viability dye. Viability dyes should be included in all compensation, control, and sort tubes.

31. To remove any cell clumps which may block the flow cytometer, aliquot samples into 5-ml FACS tubes with cell strainer caps (35-μm, sterile). To filter larger volumes of cells for sorting, a 40-μm nylon-mesh cell strainer can be used.

Set up for FACS Detailed protocols for flow cytometry are provided in Robinson et al. (2010). 32. Prepare collection tubes containing PBS with 2% fetal bovine serum. Cells can be collected into microcentrifuge tubes (containing 200 μl of PBS/2% fetal bovine serum) or 5 ml FACS tubes (containing 1 ml of the PBS/serum).

33. Set up flow cytometer with a large (90 to 100-μm) nozzle and stabilize the flow stream under low pressure (30 to 40 psi). 34. Set the compensation settings using single-color control tubes. When setting the compensation using single-color control tubes containing lung cells stained with the appropriate antibodies and fluorochromes, it is important to take into consideration the autofluorescence of lung cells and avoid overcompensation (see Critical Parameters).

35. To sort lung EpiSPC and MSC, set sequential gates for selection of viable cells (Fig. 2G.1.1B) followed by exclusion of hematopoietic (CD45) and endothelial (CD31) cells (Fig. 2G.1.1C), prior to selection of EpCAMhi CD24low EpiSPC (Fig. 2G.1.1D) and/or EpCAMneg Sca-1hi MSC (Fig. 2G.1.1E). Isotype controls should be used to account for non-specific antibody staining in setting gates to identify and isolate antibody-positive cells.

36. Sort cells into collection tubes containing PBS containing 2% fetal bovine serum.

ORGANOTYPIC CULTURE OF LUNG EPITHELIAL STEM/PROGENITOR CELLS (DIFFERENTIATION CULTURE)

BASIC PROTOCOL 2

This protocol is used for the identification and characterization of EpiSPC. During culture, epithelial CFU (Epi-CFU) proliferate and differentiate to form complex lineage-restricted or multipotent epithelial colonies comprising alveolar cells, airway cells, or cells of mixed lineage. The growth of these colonies requires seeding EpiSPC in a three-dimensional extracellular matrix (Matrigel) in co-culture with lung-derived MSC (Fig. 2G.1.2A,B). These cultures can be re-fed and maintained over 2 weeks, after which cells begin to die and colonies deteriorate. This culture technique has been developed for EpiSPC sorted on the phenotype CD45neg CD31neg EpCAMhi CD24low , but can also be used to identify EpiSPC from heterogeneous cell fractions. MSC are sorted on the phenotype of CD45neg CD31neg EpCAMneg Sca-1hi .

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Figure 2G.1.2 (A) Schematic representation of organotypic culture setup for EpiSPC differentiation. (B) Image of 24-well tissue culture plate with culture inserts. (C) Whole-well image of colonies after 2 weeks in culture. (D) Bright-field and (E) dark-field images depicting representative colony subtypes, including (i) airway, (ii) alveolar, and (iii) mixed colonies.

NOTE: All procedures are performed in a sterile class II biological hazard flow hood or a laminar-flow hood. All solutions, reagents, media and equipment used to process and culture EpiSPC must be sterile, and proper aseptic technique should be used.

Materials Epi-CFU medium (see recipe) Matrigel (standard concentration; BD Biosciences) Adult lung EpiSPC (CD45neg CD31neg EpCAMhi CD24low ) cells (Basic Protocol 1) Adult lung MSC (CD45neg CD31neg EpCAMneg Sca-1hi ) cells (Basic Protocol 1) Refrigerated centrifuge Millicell-CM culture inserts (0.4-μm membrane, 12-mm diameter, hydrophilic PTFE, Millipore) 24-well flat-bottom tissue culture plates Triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ; see Critical Parameters), humidified Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells


Additional reagents and equipment for determining viable cell concentration (UNIT 1C.3) Prepare organotypic cultures of lung EpiSPC 1. Following flow cytometric cell sorting, wash the sorted EpiSPC and MSC separately in 5 ml/tube Epi-CFU medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. Current Protocols in Stem Cell Biology

2. Resuspend individual cell pellets in Epi-CFU medium (1 to 2 ml per lung) and take an aliquot to determine cell concentration (UNIT 1C.3). Cell counts can be performed using an automated cell counter or a hemacytometer (UNIT 1C.3).

3. After determining cell concentrations, take aliquots of EpiSPC and MSC and combine so that the final mixed cell suspension contains the desired number of cells for culture. Allow for 100 μl per well. For optimal Epi-CFU growth supporting capacity, MSC (CD45neg CD31neg EpCAMneg Sca1hi ) should be used at 2 × 106 cells/ml. The concentration of sorted cells seeded for detection of EpiSPC depends on their level of enrichment in the sorted fraction. Ideally, cells should be seeded at a concentration which will generate about 20 colonies per Millicell insert (∼500 cells). The colony-forming efficiency of CD45neg CD31neg EpCAMhi CD24low EpiSPC from adult (8- to 12-week old) C57Bl/6 mice is typically 1 in 23.

4. Centrifuge EpiSPC/MSC mix 5 min at 400 × g, 4◦ C, and aspirate medium.

Prepare Matrigel suspension cultures 5. Resuspend cell pellet in Matrigel diluted 1:1 with Epi-CFU medium. Allow 100 μl per well. Ensure matrix always stays on ice; otherwise, it will solidify.

6. Gently mix the Matrigel cell suspension. It is important to avoid creating bubbles. This can be achieved by holding the tube in the center of a Vortex mixer to create a swirling motion.

7. Place 12-mm Millicell-CM inserts in 24-well culture plates. 8. Add 90 μl of Matrigel cell suspension on top of the filter membrane of a Millicell-CM insert and place plate in incubator (37◦ C) for 5 min to allow matrix to set. Be careful to avoid creating bubbles in the suspension.

9. Remove plate from incubator and add 400 μl of Epi-CFU medium per well around the insert. 400 μl is just enough to allow the medium to touch the bottom of the insert, allowing diffusion of medium into the Matrigel without submerging the Matrigel culture. This semi-dry state of the Matrigel is essential for epithelial colony-formation.

10. Incubate cell cultures in a humidified 37◦ C triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ) and change to fresh Epi-CFU medium three times weekly. 11. Visualize colony morphology by bright- or dark-field microscopy (Fig. 2G.1.2C-E). The fractionation strategy described in Basic Protocol 1 enriches for a population of cells containing both lineage-restricted airway and alveolar CFU and mixed-lineage CFU, which can be identified based on colony morphology (Fig. 2G.1.2D,E).

EXPANSION OF LUNG EPITHELIAL STEM/PROGENITOR CELLS IN CULTURE

BASIC PROTOCOL 3

In this protocol, EpiSPC are seeded in a stromal-free Matrigel culture supplemented with FGF-10 and HGF (Fig. 2G.1.3), in which they generate spherical cystic colonies that can be enzymatically dissociated and passaged weekly to maintain EpiSPC. This assay can also be used to assess the self-renewal capacity of putative EpiSPC. Somatic Stem Cells


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Figure 2G.1.3

Schematic representation of stromal-free culture setup for EpiSPC expansion.

NOTE: All procedures are performed in a sterile class II biological hazard flow hood or a laminar-flow hood. All solutions, reagents, media and equipment used to process and culture EpiSPC must be sterile, and proper aseptic technique should be used.

Materials Epi-CFU expansion medium (see recipe) Matrigel (standard concentration; BD Biosciences) Adult lung EpiSPC (CD45neg CD31neg EpCAMhi CD24low ) cells (Basic Protocol 1) Phosphate buffered saline (PBS; Sigma, cat. no. P-3813) Enzymatic digestion cocktail (see recipe) Refrigerated centrifuge Millicell-CM culture inserts (0.4-μm membrane, 12-mm diameter, hydrophilic PTFE, Millipore) 24-well flat-bottom tissue culture plates Triple-mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ), humidified 15-ml conical tubes (e.g., BD Falcon) and 2-ml microcentrifuge tubes 21-G needles Set up expansion culture of lung EpiSPC 1. Following flow cytometric cell sorting, wash the sorted EpiSPC and MSC in 5 ml/tube Epi-CFU medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. 2. Resuspend individual cell pellets in Epi-CFU medium (1 to 2 ml per lung) and take an aliquot to determine cell concentration (UNIT 1C.3). Cell counts can be performed using an automated cell counter or a hemacytometer (UNIT 1C.3).

3. After determining cell concentrations, take an aliquot of Epi-SPC so that the final suspension contains the desired number of cells for culture (1000 cells per well). Allow for 100 μl per well. 4. Centrifuge EpiSPC cell suspension 5 min at 400 × g, 4◦ C, and aspirate medium. 5. Resuspend cell pellet in Matrigel diluted 1:1 with Epi-CFU expansion medium. Allow 100 μl per well. Ensure matrix always stays on ice otherwise it will solidify. Isolation and Clonal Assay of Adult Lung Epithelial Stem/Progenitor Cells

6. Gently mix the Matrigel cell suspension. It is important to avoid creating bubbles. This can be achieved by holding the tube in the center of a Vortex mixer to create a swirling motion.

7. Place 12-mm Millicell-CM inserts in 24-well culture plates.



8. Add 90 μl of Matrigel cell suspension atop of the filter membrane of a Millicell-CM insert and place plate in incubator (37◦ C) for 5 min to allow matrix to set. Be careful to avoid creating bubbles in the suspension.

9. Remove plate from incubator and add 400 μl of Epi-CFU expansion medium per well around the insert. 400 μl is just enough to allow the medium to touch the bottom of the insert, allowing diffusion of medium into the Matrigel without submerging the Matrigel culture.

10. Incubate cell cultures in a humidified 37◦ C triple–mix incubator (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ) and change to fresh Epi-CFU expansion medium three times weekly.

Passage cells 11. After 1 week in culture, aspirate medium and wash cultures twice, each time with 1 ml sterile PBS. It is important to remove serum-supplemented medium because the serum will inhibit subsequent enzymatic digestion.

12. Harvest epithelial CFU by adding 1 ml of enzymatic digestion cocktail to the top of the insert and break up Matrigel by trituration. If clonal passaging is required, single colonies can be picked from the Matrigel and enzymatically digested rather than the bulk culture.

13. After Matrigel has been displaced from the insert, place enzymatic digestion cocktail (containing Matrigel and colonies) in a 2-ml microcentrifuge tube and incubate at 37◦ C for 30 min. 14. Using a 21-G needle, triturate the enzymatic digest to prepare a single-cell suspension. 15. Wash twice, each time with 5 ml Epi-CFU expansion medium, centrifuge 5 min at 400 × g, 4◦ C, and aspirate medium. 16. Re-seed single-cell suspension by repeating steps 5 to 10. Cells should be passaged weekly and can be split into multiple cultures to accommodate the increase in cell concentration. The number of cells re-seeded depends on the progressive enrichment of colony-forming cells after sequential passage.


Enzymatic digestion cocktail PBS (tissue-culture grade; Invitrogen, cat. no. 14040) containing: 3 mg/ml collagenase (Type I) 3 mg/ml dispase Preheat at 37◦ C and use immediately Epi-CFU expansion medium Epi-CFU medium (see recipe) containing: 50 ng/ml recombinant FGF-10 (R&D Systems, cat. no. 345-FG) 30 ng/ml recombinant HGF (R&D Systems, cat. no. 2207-HG) Store up to 1 week at 4◦ C

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Epi-CFU medium α-MEM (Invitrogen, cat. no. 41061) containing: 10% (v/v) fetal bovine serum (heat inactivated) 1× penicillin/streptomycin (add from 100× stock) 1× insulin/transferrin/selenium (Invitrogen; add from 100× stock) 2 mM L-glutamine 0.0002% (w/v) heparin [1/1000 dilution of 0.2% heparin sodium salt (Invitrogen, cat. no. 07980) in PBS (Invitrogen, cat. no. 14040)] Store up to 4 weeks at 4◦ C Nycoprep 1.077 g/cm3 , 265 mOsm Combine the following: 300 ml Nycoprep Universal: (Nycodenz: 60% w/v solution), ready-made, sterile, endotoxin-tested, density = 1.310 g/cm3 ; 580 mOsm; 300 ml (Axis-Shield; http://www.axis-shield.com/)

300 ml sterile Tricine-NaOH (20 mM, pH = 7.2) 676.6 ml sterile 0.65% NaCl (w/v) Density = 1.077 gm/cm3 , Osmolarity = 265 mOsm, pH = 6.9 Store at room temperature and use before manufacturer’s expiration date COMMENTARY Background Information


Identification and characterization of adult lung EpiSPC have been confounded by a lack of specific markers and functional assays for their prospective isolation, enumeration, and measurement of their proliferative and differentiative potential (Weiss et al., 2008; Chen et al., 2009; Bertoncello and McQualter, 2010). A number of studies have utilized flow cytometry for isolation of candidate lung stem/progenitor cell populations, including those based on the efflux of Hoechst 33342 which has proven to be selective for enriching Sca-1pos mesenchymal stromal cells (MSC; Reynolds et al., 2007; Summer et al., 2007). We have also demonstrated that sorting directly on the basis of Sca-1 expression enriches for MSC (McQualter et al., 2009). On the other hand, Kim et al. (2005) have reported a fractionation strategy in which sorting based on the co-expression of Sca-1 and CD34 resulted in the enrichment of a candidate CCSPpos Pro-SPCpos bronchioalveolar stem cell (BASC) cell subpopulation that retained epithelial character after serial passage in vitro. This protocol describes a strategy for isolating CD45neg CD31neg EpCAMhi CD24low lung EpiSPC, which are Sca-1low and clearly distinct from CD45neg CD31neg EpCAMneg Sca-1hi MSC or BASC (McQualter et al., 2010). The lack of concordance in the properties of cells isolated in these studies could

be explained by technical differences in tissue disaggregation (Raiser and Kim, 2009) and the limitations of in vitro assays used to assess proliferative and differentiative potential. The lack of knowledge of the intricate interactions between epithelial cells, mesenchymal cells, and the extracellular matrix has proven a significant obstacle in recapitulating the necessary conditions in vitro required for the development of assays for the identification of EpiSPC and the analysis of their organization and regulation. The culture system described in this unit utilizes a three-dimensional extracellular matrix (Matrigel), which allows the formation of a basement membrane for epithelial cell polarization and lumen formation, and enables the organotypic rearrangement of cells in culture recapitulating the physiological microenvironment of the lung. For that reason, this culture system can also be used to study epithelial-mesenchymal interactions that are important for lung regeneration and repair.

Critical Parameters To correct for spectral overlap between different fluorochromes during multicolor flow cytometric analysis and sorting, color compensation must be performed to correctly quantify the fluorescence intensity of each fluorochrome with which cells are labeled. When setting the level of compensation using cells from dissociated lung tissue, it is important to



Figure 2G.1.4 Bivariate dot plot with back gating of EpiSPC (red) and MSC (blue) onto CD45neg CD31neg cells (gray) showing the potential overlap of these populations when plotted as CD45 + CD31 versus Sca-1.

take into account the autofluorescence of these cells. Alignment of autofluorescent cells with nonautofluorescent cells will result in overor under-compensation and inaccurate assignment of cell phenotype. Correct compensation assumes that the positive and negative populations have equal autofluorescence (Alexander et al., 2009; Alvarez et al., 2010). When applying the gating strategy used to sort cells by flow cytometry it is important to understand that very rarely are the boundaries between populations absolute. We have previously isolated lung MSC based on the CD45neg CD31neg Sca-1pos phenotype, which also comprised a minor population of epithelial CFU (McQualter, et al., 2009). However, subsequent introduction of EpCAM to the sort strategy demonstrated that all epithelial CFU could be removed from the MSC fraction by gating on the CD45neg CD31neg EpCAMneg Sca-1hi phenotype. Figure 2G.1.4 shows that when EpiSPC and MSC are back-gated onto our traditional plot of CD45+CD31 versus Sca1, the two subsets marginally overlap, which would explain why gates previously set for MSC using the CD45neg CD31neg Sca-1pos phenotype may also include a minor fraction of EpiSPC (McQualter et al., 2010). This protocol describes a method in which EpiSPC are cultured within Matrigel atop a Millicell-CM insert with medium supplied only at the basal surface to allow diffusion of nutrients into the Matrigel culture. It is important that the Matrigel layer not be submerged by medium, as this prevents colony formation. The Millicell-CM inserts chosen for this assay contain a special Biopore membrane (hydrophilic PTFE), which helps limit overgrowth of the stromal layer, and is transparent, allowing microscopic visualization.

The Epi-CFU described in these protocols have been grown under physiological low oxygen tension (5% v/v O2 , 10% v/v CO2 , 85% v/v N2 ), which has been shown to be optimal for growth of stem/progenitor cells at clonal density in vitro (Wion et al., 2009). However, Epi-CFU can be grown under standard oxygen tension (10% v/v CO2 in air), but the cloning efficiency may be lower.

Troubleshooting It is the authors’ experience that the differentiation state of MSC used in organotypic co-cultures is critical for supporting the growth of EpiSPC. It is important to use fresh CD45neg CD31neg EpCAMneg Sca-1hi MSC, as expansion of these cells in culture results in their differentiation and inhibits their ability to support growth of EpiSPC (manuscript in preparation).

Anticipated Results Using this protocol, the cell CD45neg CD31neg EpCAMhi CD24low fraction isolated comprises an enriched but heterogeneous population of lineagerestricted (airway or alveolar) epithelial progenitors and multipotent (multi-lineage) stem cells (∼2000 cells per lung), while the CD45neg CD31neg EpCAMneg Sca-1hi cell fraction represents a population of enriched MSC (∼100,000 cells per lung; McQualter et al., 2010). EpiSPC are cultured using two different techniques. Morphological characterization of colonies generated from EpiSPC grown in organotypic differentiation cultures demonstrates the generation of large lobular cystic colonies with a clearly defined lumen (airway-CFU), small dense saccular colonies

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(alveolar-CFU), and colonies of mixed phenotype with distinct budding (multipotentCFU, Fig. 2G.1.2). Under these conditions, MSC differentiate into lipid-filled fibroblasts and myofibroblasts. In stromal-free expansion cultures, EpiSPC generate smaller spherical colonies, which can be enzymatically passaged on a weekly basis.

Time Considerations Temporal analysis of colony formation in this organotypic assay system (Basic Protocol 2) results in the emergence of colonies after 5 days in culture, and their continued expansion and differentiation over a 2-week period. In stromal-free expansion cultures (Basic Protocol 3), optimal colony formation for serial propagation and re-seeding of CFU is achieved when colonies are harvested after 1 week. After this point, colonies begin differentiate and deteriorate, and the recloning efficiency of dissociated colonies is substantially reduced.

Literature Cited Alexander, C.M., Puchalski, J., Klos, K.S., Badders, N., Ailles, L., Kim, C.F., Dirks, P., and Smalley, M.J. 2009. Separating stem cells by flow cytometry: Reducing variability for solid tissues. Cell Stem Cell 5:579-583.

Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Kim, C.F., Jackson, E.L., Woolfenden, A.E., Lawrence, S., Babar, I., Vogel, S., Crowley, D., Bronson, R.T., and Jacks, T. 2005. Identification of bronchioalveolar stem cells in normal lung and lung cancer. Cell 121:823-835. McQualter, J.L., Brouard, N., Williams, B., Baird, B.N., Sims-Lucas, S., Yuen, K., Nilsson, S.K., Simmons, P.J., and Bertoncello, I. 2009. Endogenous fibroblastic progenitor cells in the adult mouse lung are highly enriched in the sca-1 positive cell fraction. Stem Cells 27:623633. McQualter, J.L., Yuen, K., Williams, B., and Bertoncello, I. 2010. Evidence of an epithelial stem/progenitor cell hierarchy in the adult mouse lung. Proc. Natl. Acad. Sci. U.S.A. 167:1414-1419. Raiser, D.M. and Kim, C.F. 2009. Commentary: Sca-1 and cells of the lung: A matter of different sorts. Stem Cells 27:606-611. Reynolds, S.D., Shen, H., Reynolds, P.R., Betsuyaku, T., Pilewski, J.M., Gambelli, F., Di Giuseppe, M., Ortiz, L.A., and Stripp, B.R. 2007. Molecular and functional properties of lung SP cells. Am. J. Physiol. Lung Cell Mol. Physiol. 292:L972-L983. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P, Orfao, A., Rabinovitch, P., and Watkins, S. 2010. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J.

Alvarez, D.F., Helm, K., Degregori, J., Roederer, M., and Majka, S. 2010. Publishing flow cytometry data. Am. J. Physiol. Lung Cell Mol. Physiol. 298:L127-L130.

Summer, R., Fitzsimmons, K., Dwyer, D., Murphy, J., and Fine, A. 2007. Isolation of an adult mouse lung mesenchymal progenitor cell population. Am. J. Respir. Cell Mol. Biol. 37:152159.

Bertoncello, I. and McQualter, J.L. 2010. Endogenous lung stem cells: What is their potential for use in regenerative medicine? Expert Rev. Respir. Med. 4:349-362.

Weiss, D.J., Kolls, J.K., Ortiz, L.A., PanoskaltsisMortari, A., and Prockop, D.J. 2008. Stem cells and cell therapies in lung biology and lung diseases. Proc. Am. Thorac. Soc. 5:637-667.

Chen, H., Matsumoto, K., and Stripp, B.R. 2009. Bronchiolar progenitor cells. Proc. Am. Thorac. Soc. 6:602-606.

Wion, D., Christen, T., Barbier, E.L., and Coles, J.A. 2009. PO(2) matters in stem cell culture. Cell Stem Cell 5:242-243.




Colon Cancer Stem Cells 1

UNIT 3.1 2, 3

Antonija Kreso and Catherine Adell O’Brien 1

Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada 2 Institute of Medical Sciences, University of Toronto, Toronto, Ontario, Canada 3 Department of Surgery, Division of General Surgery, University Health Network, Toronto, Ontario, Canada

ABSTRACT This unit describes protocols for working with colon cancer stem cells. To work with these cells one must start by generating single-cell suspensions from human colon cancer tissue. These cell suspensions are sorted using flow cytometry–assisted cell sorting to fractionate the cells into tumor-initiating and nontumor-initiating subsets. Once the cells have been fractionated, they must be functionally tested to determine tumor-forming capacity, the gold standard being the in vivo xenograft assay. Methods have also been developed to grow these cells in vitro in a sphere-forming assay. This unit will describe how to isolate and functionally test colon cancer stem cells, as well as provide advice on the potential challenges of the research. Curr. Protoc. Stem Cell Biol. 7:3.1.1-3.1.12. C 2008 by John Wiley & Sons, Inc. Keywords: human colon cancer r cancer stem cells r in vivo xenograft assay r in vitro sphere assay

INTRODUCTION This unit describes protocols for working with colon cancer stem cells (CSC). The ability to successfully carry out this work is dependent on obtaining fresh colon cancer specimens at the time of surgical resection. Tissue fragments are processed to generate a single-cell suspension, which can then be fractionated utilizing flow cytometry to isolate subpopulations based on differential expression of cell surface markers, such as CD133 (O’Brien et al., 2007; Ricci-Vitiani et al., 2007; Todaro et al., 2007). Once these cell subsets have been fractionated, they can be tested for their tumor-forming capacity using the in vivo NOD/SCID xenograft assay. Utilizing this model it has been shown that tumor-initiating capacity exists solely within the CD133+ cell subset of colon cancer cells. The focus of this unit will be to describe the protocols for isolating, culturing (Basic Protocol 1), fractionating (Basic Protocol 2), and establishing a NOD/SCID xenograft model (Basic Protocol 3) to study colon CSC. The sphere-forming assay is also described (Basic Protocol 4). NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator. NOTE: All experiments using human tissue must be approved by the institutional committee on the ethical use of human subjects/material and tissue samples must be obtained with prior informed consent. Cancer Stem Cells Current Protocols in Stem Cell Biology 3.1.1-3.1.12 Published online November 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0301s7 C 2008 John Wiley & Sons, Inc. Copyright

3.1.1 Supplement 7

NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals. BASIC PROTOCOL 1

GENERATING SINGLE-CELL SUSPENSIONS FROM HUMAN COLON CANCER TISSUE The first step to working with colon CSC involves generating a single-cell suspension from human colon cancer tissue. The percentage of necrotic cells in human tumors varies extensively and is dependent on multiple factors including: tumor characteristics, preoperative adjuvant chemotherapy or radiation therapy, and length of operative procedure. One factor that can help in obtaining the maximum number of viable cells is to ensure that specimens are received from the operating room expeditiously after removal from the patient. It has been our experience that each sample possesses a CSC fraction; however, the percent of this fraction can vary widely between tumors and this is true whether one is using CD133 or CD44 to isolate the CSCs. It is best when isolating the CSC fraction to start with at least 1 to 2 × 106 colon cancer cells (use a tumor fragment ∼1 × 0.5–cm in size to generate this many cells), because this will help ensure that there are enough cells in the CSC and non-CSC subsets to carry out the experiments.

Materials Colon tumor fragment Colon cancer stem cell medium (SCM; see recipe) Collagenase IV solution (200 U/ml SCM) Ammonium chloride: 0.8% (w/v) NH4 Cl in 0.1 mM EDTA Trypan blue 35-mm petri dishes Razor blade and forceps 5-ml disposable pipets 50-ml conical tube 45-μm cell filter Plunger from a 3- to 5-ml syringe Hemacytometer Additional reagents and equipment for counting cells using a hemacytometer and trypan blue (UNIT 1C.3) Isolate colon cancer cells 1. Place colon tumor fragment in 2 to 3 ml SCM in a 35-mm petri dish. 2. Using a razor blade and forceps, mince the tissue as much as possible. 3. Pipet tumor solution up and down 3 to 5 min with a 5-ml disposable pipet. Place the solution into a 50-ml conical tube. If fragments are too large to be drawn up into a 5-ml pipet, use a pipet with an opening large enough to draw up all the tumor fragments. Note that the highest cell numbers are typically obtained when tumors are minced to yield very small pieces.

4. Add the collagenase solution to the tumor cells. Incubate 30 to 60 min at 37◦ C. Pipet up and down a few times every 15 min. The final concentration should be 200 U of collagenase IV per milliliter of SCM.

Colon Cancer Stem Cells

5. Pass the tumor solution through a 45-μm filter. Use a plunger from a 3- to 5-ml syringe and gently mash the tumor pieces to enable more tumor cells to pass through. Wash the filter with 4 to 5 ml of SCM.

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The tumors can be very different; some are soft whereas others are hard and fibrotic. The fibrotic tumors may not completely dissolve with collagenase. The fragments that remain on the filter can be resuspended in SCM with collagenase and incubated for 1 to 2 hr at 37o C. Then repeat step 5.

6. Centrifuge the tumor cell suspension 10 min at 450 × g, 4◦ C. 7. Resuspend the pellet in ∼5 ml of ammonium chloride (0.8% w/v NH4 Cl with 0.1 mM EDTA). Leave for 10 min at room temperature to lyse the red blood cells. After 10 min add an equal volume of SCM and centrifuge 10 min at 450 × g, 4◦ C. 8. Resuspend the pellet in 10 ml SCM. If the solution appears clumpy then pass it through another 45-μm filter. 9. Count an aliquot of the cells using a hemacytometer and trypan blue (UNIT 1C.3) to determine the percentage of dead cells. If there is a high percentage of necrotic cells, a Ficoll column can be used to remove the dead cells and debris (see Support Protocol).

USING A FICOLL COLUMN TO REMOVE DEAD CELLS The high percentage of necrotic cells and debris in some samples makes it exceedingly difficult to successfully carry out techniques such as flow cytometry–assisted cell sorting and transduction. In cases where samples have >30% dead cells this Ficoll protocol can allow for an enrichment of viable cells.

SUPPORT PROTOCOL

Materials Ficoll Colon cancer cell suspension (Basic Protocol 1) Colon cancer stem cell medium (SCM; see recipe) 15-ml conical tubes 5-ml pipet 1. Place 5 ml Ficoll into a 15-ml conical tube. 2. Resuspend the colon cancer cells in 5 ml of SCM. Layer this solution on top of the 5 ml of Ficoll. Divide the tumor cell suspension such that each 5 ml of medium contains no more than 4 to 5 × 106 tumor cells. If the cell number is >5 × 106 , divide the sample into the appropriate number of Ficoll-containing tubes.

3. Centrifuge 15 min at 1000 × g, 4◦ C. 4. Use a 5-ml pipet to remove 2 to 3 ml of medium off the top and then place the pipet at the interface (between the medium and Ficoll). Collect the interface, remainder of the medium, and a small amount of Ficoll. Resuspend the pellet in 5 ml SCM and save until the viable cell count is complete. Keep this solution until the cell number from the viable fraction has been counted. If the number of viable cells post-Ficoll differs significantly from the pre-Ficoll count, it is possible that some viable cells are in the pellet. In that case, repeat step 2 with the resuspended pellet.

5. Centrifuge 10 min at 450 × g, 4◦ C. Resuspend the pellet in a desired volume of SCM. Cancer Stem Cells

3.1.3 Current Protocols in Stem Cell Biology

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BASIC PROTOCOL 2

FLOW CYTOMETRY–ASSISTED CELL SORTING Flow cytometric–assisted cell sorting is an essential aspect of CSC work. It is required to fractionate the CD133+ and CD133− cell subsets. It also allows the researcher to exclude all murine cells when sorting colon cancer xenografts, thereby avoiding any murine hematopoietic or endothelial cells contaminating the post-sort cell populations. To avoid contaminating cells in primary human colon cancer sorts one can positively select for epithelial specific antigen (ESA) expression and sort the following populations: ESA+ CD133+ and ESA+ CD133− . The initial selection on ESA+ cells allows one to exclude contaminating hematopoietic, endothelial, and stromal cells. Furthermore, flow cytometry also allows one to study other markers of interest in combination with CD133. If flow cytometry–assisted cell sorting is not available, another option is to carry out magnetic bead cell sorting as per the Miltenyi-Biotec protocol. It has been our experience that this method can be used successfully provided the starting sample has ≤30% dead cells. If the sample has >30% dead cells, it can be difficult to obtain the necessary purity (≥90% to 95%) using the MACS bead separation. To successfully carry out MACS bead enrichment the sample should be passed through at least three Miltenyi-Biotec columns in order to best enrich the sample. One of the other main disadvantages of using the MACS beads is the inability to select on multiple cell surface markers.

Materials Colon cancer cell suspension with 5 × 106 , divide the cells into separate polystyrene tubes for staining. This is done because it is best to stain cells in a smaller volume.

2. Add fluorophore-conjugated anti-CD133 and anti-mouse or anti-ESA antibodies to the sort sample. When sorting human samples, an antibody against ESA should be used. When sorting xenografts, use an anti-murine IgG antibody to exclude murine cells. Check with the FACS facility to determine whether isotypes and individual antibody control stains are required. All antibodies should be titrated to determine the required amount. Ask the FACS facility whether they prefer that the sort sample be placed into a polypropylene or polystyrene tube for sorting.

3. Incubate cells with antibodies 30 to 45 min at 4◦ C, protected from light. 4. Add 4.5 ml CMF-PBS/0.1% BSA to each tube and centrifuge 5 min at 450 × g, 4◦ C. Discard the supernatant. Repeat three times to wash the cells. Colon Cancer Stem Cells

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5. Following the last wash, resuspend the cells in 1 to 2 ml CMF-PBS/0.1% BSA that contains 1 μg/ml propidium iodide. At this point the cells are ready to be sorted. Propidium iodide stains dead cells thereby allowing them to be excluded.

6. Sort the ESA+ CD133+ cells (Robinson et al., 2008) into 2 ml SCM. 7. Count an aliquot of the post-sort cells with a hemacytometer (UNIT 1C.3) to confirm the cell yield. The flow cytometry facility will provide a post-sort cell count for each population; however, it is important that the researcher confirm the cell count.

IN VIVO XENOGRAFT ASSAY The utilization of animal models is crucial in CSC work. The ideal choice is always an orthotopic model in which cancer cells are injected into the same tissue from which they are derived. However, when carrying out CSC research the most important factor is to identify an animal model that has the greatest reliability for xenograft formation. This is essential because if the tumor take rate for the model you choose is only 50% it becomes impossible to determine whether the absence of a xenograft is due to the lack of a CSC or simply the limitation of the animal model. Published colon CSC work to date have used two models: subcutaneous (Ricci-Vitiani et al., 2007; Todaro et al., 2007) and subrenal capsule (O’Brien et al., 2007) injections. It has been our experience that only ∼30% of colon cancer cell suspensions have a reliable take rate in the subcutaneous site. In our hands, injection of colon cancer cell suspensions under the renal capsule provided the most reliable results with almost all tumors tested giving rise to xenograft formation (overall take rate of ∼90%).

BASIC PROTOCOL 3

Aside from the choice of injection site a decision must also be made about the type of immunocompromised mouse to be used. Published studies in colon CSC research have used either NOD/SCID or SCID mice (Dalerba et al., 2007; O’Brien et al., 2007; RicciVitiani et al., 2007). It is difficult to make a direct comparison between the efficiency of the NOD/SCID versus SCID mice for use in CSC work because no head-to-head comparison has been carried out between the two strains using the same marker set and injection site. It is also important to acknowledge that an increasing number of immunocompromised mouse strains are becoming available and may represent new options for carrying out this work. Irradiating the mice prior to the procedure can also improve the xenograft take rate; however, the marginal improvement in xenograft formation must be weighed against the radiation sensitivity of NOD/SCID mice. Irradiation should be carried out the day before or the day of the procedure and the dose should be 300 rad. It is best to carry out a trial of irradiation to determine both the potential benefit to xenograft take rate and the radiation sensitivity of the mice in the colony. Death related to radiation sensitivity usually occurs 6 to 7 weeks post-irradiation.

Materials Matrigel ∼10 μl sorted cell suspension in SCM (Basic Protocol 2) Stem cell medium (SCM; see recipe) NOD/SCID or SCID mice (8- to 10-week-old) Iodine-based solution (e.g., Betadine) 70% ethanol

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Normal saline, sterile Pain medication (e.g., buprenorphine) 1-ml syringe without needle 1-ml insulin syringes with a 29-G needle, 1/2 -in. long Heating pad Clippers Sterile gauze Scissors Forceps Sutures or surgical clips (Roboz) Additional reagents and equipment for rodent anesthesia using isoflurane (UNIT 1B.4) 1. Thaw an aliquot of Matrigel and draw it up into a 1-ml syringe without needle. Then remove the plunger from a 1-ml insulin syringe with a 29-G needle, 1/2 -in. long and insert a small amount of Matrigel into the back of the syringe (Fig. 3.1.1A). It is difficult to be exact on the amount of Matrigel—one should aim for it to be 25 to 50 μl (closer to 25 μl is better). The best way to estimate the amount of Matrigel is to look at the markings on the side of the syringe. Keep undiluted Matrigel aliquoted and frozen at −20o C, ∼100 to 200 μl per microcentrifuge tube.

2. Use a pipet to inject the cell suspension (aim to resuspend the cells in 10 μl of SCM) into the middle of the Matrigel (Fig. 3.1.1B). Then reinsert the plunger into the syringe and push the mixture to the top of the syringe (Fig. 3.1.1C,D). Once the syringes are prepared they should be injected in a timely fashion. It is best to keep the syringes on ice, because at room temperature Matrigel will set.

Figure 3.1.1 Preparing syringes for injection. (A) Inserting the Matrigel into the back of the insulin syringe. (B) Using a pipet place the 10 μl of cell solution into the middle of the Matrigel. (C) Reinsert the insulin syringe plunger. (D) Using the plunger push the Matrigel and cell solution to the top of the syringe, it is now ready for injection. Colon Cancer Stem Cells

3.1.6 Supplement 7


Figure 3.1.2 Subrenal capsule injection. (A) The mouse is positioned with its left side up. A left flank incision is made just under the costal margin. (B) The kidney is gently pulled out of the abdominal cavity, being careful not to disturb the blood supply. (C) The needle is positioned just under the renal capsule. (D) Following the injection of Matrigel and cell suspension there is a small bleb on the surface of the kidney.

3. Handle mice using sterile technique and anesthetize using inhalational anesthesia (UNIT 1B.4). Place the mice on a heating pad during the procedure. 4. Position the mouse left side up. Use clippers to shave the area ∼0.5-cm below the costal margin on the left side. Wash the clipped area sequentially with iodine-based solution and 70% ethanol solution and then dab dry with sterile gauze. 5. Using scissors make an ∼0.5-cm long incision on the flank, just below the costal margin on the left side (Fig. 3.1.2A). Only inject into the left kidney. Due to the anatomy it is very difficult to inject into the right kidney. Cancer Stem Cells


Supplement 7

6. Gently pull the kidney from the abdominal cavity using two pairs of forceps. Then take the syringe containing the cells to be injected and place the needle just under the kidney capsule and push the needle until just before it comes out the opposite pole. At this point, inject the cells as you slowly pull back on the needle. The cell solution should be completely injected before the needle exits the kidney (Fig. 3.1.2B,C,D).

7. Deliver the kidney back into the abdomen and close the abdominal wall. There is the choice to close using sutures or surgical clips (Roboz) both of which provide equivalent results. The choice may depend on the requirements of the animal research facility at your particular institution.

8. Prior to awakening the mouse from anesthesia administer a 1 ml subcutaneous bolus of sterile normal saline and a dose of pain medication (e.g., 0.01 to 0.05 mg/kg buprenorphine). At the commencement of this work an antibiotic (Baytril) was added to the drinking water of all mice post-procedure for 2 weeks. The mice became dehydrated and as a result weak; therefore, the practice was stopped. There were no deleterious effects from discontinuing Baytril.

9. Assess the mice for tumor development every week starting 2 weeks post procedure. This can be done by holding the mouse and gently palpating in the area of the kidney. Over time, you will start to appreciate a fullness (the tumor will feel like a firm nodule) in this area. The rate of tumor development will differ depending on the tumor. Some xenografts will appear in 6 to 8 weeks; however, others can take up to 30 weeks to develop. BASIC PROTOCOL 4

CULTURING COLON CANCER CELLS AS SPHERES The ability to culture colon CSC requires the utilization of a serum-free stem cell medium. Using this protocol colon CSC will grow as spheres (Fig. 3.1.3A,B) in a non-adherent manner. The addition of serum to this medium results in the differentiation of the colon cancer cells and their growth as an adherent layer. Although much of the current work in the cancer stem cell field has been carried out using in vivo models, the ability to culture the cells as spheres can be used to complement the in vivo work. It is important to keep a record of the passage number for each tumor in vitro.

Materials Colon cancer cell suspension, sorted (Basic Protocol 2) Stem cell medium (SCM; see recipe) Trypsin/EDTA Ultra-low attachment surface dishes (Corning) 5-ml disposable pipet 45-μm filter Additional reagents and equipment for counting cells using trypan blue (UNIT 1C.3) 1. Plate sorted colon cancer cells at a density of 30,000 to 50,000 cells/ml of SCM. For best results the spheres should be grown in ultra-low attachment surface dishes (Corning). Any size dish can be used depending on the cell number; the most important point is to plate at a density of 30,000 to 50,000 cells/ml of SCM.

2. Passage the cells approximately every 4 days. Colon Cancer Stem Cells

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There is some variability between tumors and, therefore, each tumor must be evaluated daily to follow sphere formation. Passaging requires the disruption of the spheres to generate a single-cell suspension. There are two possible approaches to sphere disruption: mechanical or enzymatic. Current Protocols in Stem Cell Biology

Figure 3.1.3 Photographs of colon cancer sphere cultures. (A) Three colon cancer spheres (magnification 4×). (B) One colon cancer sphere (magnification 20×).

Mechanical sphere disruption 3a. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 4a. Resuspend the pellet in 3 ml of SCM and pipet up and down for 10 min with a 5-ml disposable pipet. After pipetting for 10 min look at the solution. There should no longer be any visible spheres. If visible spheres remain, continue to pipet for another 5 to 10 min.

5a. Pass solution through a 45-μm filter, count an aliquot of the cells (UNIT 1C.3), and resuspend in the desired volume of SCM for replating. Best results are obtained when cells are plated at a density of 30,000 to 50,000 cells/ml of SCM.

Enzymatic sphere disruption 3b. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 4b. Resuspend the pellet in 3 to 5 ml of 1× trypsin/EDTA and pipet up and down for 3 min with a 5-ml disposable pipet. At this point, place the tube in the incubator 10 min at 37◦ C. 5b. After 10 min remove the cells from the incubator and add an equal volume of SCM. Pass the cells through a 45-μm filter. Centrifuge the colon cancer cell solution 10 min at 450 × g, 4◦ C. 6b. Resuspend the cell solution and count the number of viable cells using trypan blue (UNIT 1C.3). The criticism associated with the use of enzymatic digestion is that it may interfere with the expression of cell surface markers. Enzymatic digestion can be used; however, it should be tested initially against mechanically digested cells from the same tumor. This testing should be carried out for each cell surface marker to determine whether enzymatic digestion has any deleterious effect on the expression of the cell surface markers being studied.

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Growth factor mix, 10× For 200 ml of growth factor mix: 100 ml DMEM/F12 4 ml 30% (w/v) glucose 200 mg transferrin 50 mg insulin in 20 ml of water (add 2 ml of 0.1 N HCl to dissolve, then add 18 ml of water) 19.33 mg putrescine in 20 ml water 200 μl 0.3 mM sodium selenite 20 μl 2 mM progesterone H2 O to 200 ml Divide the growth factor mix into 2-ml aliquots and store indefinitely at −20◦ C Stem cell medium (SCM) 500 ml of a 1:1 ratio of DMEM/F12 (Invitrogen) 1% penicillin-streptomycin (1× from a 100× stock purchased from Invitrogen) 2 ml 50× B27 supplement (Invitrogen) 4 μg/ml heparin 1% (w/v) of non-essential amino acids 1% (w/v) of sodium pyruvate 1% (w/v) of L-glutamine Store up to 1 week at 4◦ C Just before use, add 5 ml growth factor mix (see recipe) 10 ng/ml fibroblast growth factor 20 ng/ml epidermal growth factor COMMENTARY Background Information


The existence of a CSC fraction was first studied in the context of human leukemia. Lapidot et al. demonstrated that acute myelogenous leukemias possess a CSC subset capable of recapitulating the disease in a SCID mouse model, whereas the non-CSC cells were incapable of generating the disease (Lapidot et al., 1994). More recently, it has been shown that a wide variety of solid tumors also possess a CSC subset including breast (Al-Hajj et al., 2003), brain (Singh et al., 2004), and colon cancers (O’Brien et al., 2007; Ricci-Vitiani et al., 2007). The identification of a CSC population in human colon cancer was first published in 2007, when two groups established that fractionation of colon cancer cells based on CD133 expression identified a subset of CD133+ cancer cells that was capable of initiating tumor growth in murine xenograft models. In con-

trast, the CD133− cancer cells were unable to initiate tumor growth. The limiting dilution analyses in one of the studies demonstrated that ∼1 in 262 CD133+ colon cancer cells represented a CSC, for the ten tumors tested in the series, thereby demonstrating that CD133 expression enriches for tumor initiating capacity but does not identify a pure CSC population (O’Brien et al., 2007). More recently, another publication demonstrated that CD44 and CD166 (ALCAM) expression could also be utilized to enrich for a CSC subset in colon cancers (Dalerba et al., 2007). It is important to appreciate that the markers identified to date enrich for CSC; however, they do not identify a pure population. The field remains at a very nascent stage, and advancement will depend in large part on the identification of new CSC markers that can be used in conjunction with markers, such as CD133 and CD44, to provide further enrichment of the CSC fraction.

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Limiting dilution analysis (LDA) represents an essential tool in carrying out CSC work because it provides the ability to calculate the frequency of CSC within a population of cancer cells. LDAs require the injection of a range of doses with multiple mice being injected per dose (Porter and Berry, 1964). The gold standard is to carry out an LDA of both bulk and fractionated cancer cells for each individual tumor. This allows for the calculation of a CSC frequency in both the bulk tumor cell population and the fractionated subpopulations (e.g., CD133+ versus CD133− ). The limiting factor in these experiments is often the paucity of cell number. One method to circumvent issues of cell number is to initially inject unsorted cells into four to five mice to expand the cell number and then to use these xenografts to carry out the bulk and fractionated LDAs in mice. It has been our experience and the experience of others that the cell surface phenotype is maintained following passage in mice (Dalerba et al., 2007; O’Brien et al., 2007; Ricci-Vitiani et al., 2007). However, it is very important to determine the cell surface phenotype for each tumor prior to passage in mice and then after each passage. This will allow you to confirm for each tumor that the subpopulations are remaining stable following passage in vivo. It has been our experience that at high levels of in vivo passage (7 and above) we do start to see some tumors that change their cell surface phenotype; however, the changes are not predictable. It is for these reasons that it is crucial to check the tumor cell surface phenotype with each passage. There is very limited data on the propagation of colon CSC as sphere-forming units. One recent publication identified that in a series of colon cancers only approximately half could be propagated in vitro as spheres (Todaro et al., 2007). There is also the suggestion that CSC marker expression may change with in vitro propagation (A. Kreso and C.A. O’Brien, unpub. observ.). Therefore, the sphere-forming assay represents a surrogate; however, it does not eliminate the need to carry out the gold standard, functional in vivo assays. The sphere-forming assay requires further study to clearly establish its role within CSC work and to determine how closely it recapitulates in vivo models. Furthermore, caution must be exercised when using cancer cells following serial passages in vitro because to date it has not been clearly established whether the cells maintain the same functional phenotypes. Until these questions have been answered, the best approach when

using a sphere-forming assay is to functionally test the CSC and non-CSC fractions with in vivo assays at a minimum of every other in vitro passage.

Critical Parameters and Troubleshooting The ability to successfully sort the cells using flow cytometry will depend in large part on the flow facility. It is important when starting this work to determine the level of expertise at your flow facility for sorting solid tumor cells. If the facility does not have expertise in this area, it is important to contact a flow facility that regularly sorts solid tumor cells to establish the instrument settings that result in the highest yield, both with respect to purity and viability of the cells post-sorting. It is also essential to count cells post-sorting to confirm the cell yield. There can be a discrepancy between the stated and actual cell yields; having an accurate cell count is essential when carrying out LDA experiments.

Anticipated Results The in vivo protocol will result in the generation of xenografts, which recapitulate the phenotype of the original tumors. At the time of sacrifice a fragment of each xenograft should be saved for histological assessment. This will allow the researcher to confirm that the xenograft recapitulates the original tumor with respect to differentiation status and tumor subtype. The in vitro protocol will result in expansion of the colon cancer cells; however, as previously stated there is a proportion of the cancers that cannot be successfully passaged in vitro. At this time we are unable to predict which tumors will grow well in vitro. As more work is carried out in the field, it will hopefully lead to a better understanding of the in vitro requirements to maintain colon cancer cells in serum-free culture conditions.

Time Considerations The initial culturing of colon CSC from an unpassaged primary human specimen may take >4 to 5 days. If this is the case, one should leave the cells in the incubator for 7 to 14 days, adding EGF and FGF to the medium every 4 days. If the tumor has not formed spheres by 14 days it is unlikely to do so. The time for xenograft formation varies between tumors, ranging from 6 to 24 weeks. To do the in vivo experiments it is important to use NOD/SCID mice between the ages of 8 and 10 weeks. Mice utilized before 8 weeks

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have an increased chance of succumbing to the stress of the procedure. Using older mice (>10 weeks) can be difficult because as NOD/SCID mice age there is a natural attrition rate; therefore in the case of tumors that take 24 weeks to appear it is possible that the mice of interest will die before the appearance of xenografts. The primary tissue is often received from the operating room at the end of the workday and it is important to note that once the cell suspension is generated these cells can be safely left at 4◦ C overnight in SCM and injected or sorted the next day.

Acknowledgements We thank Sean McDermott for his contributions to the Ficoll protocol and other members of the Dick laboratory for helpful discussions. We also acknowledge members of Peter Dirks’ laboratory, especially Ian Clarke, for technical suggestions. We thank John E. Dick for his invaluable help and guidance.

Literature Cited Al-Hajj, M., Wicha, M.S., Benito-Hernandez, A., Morrison, S.J., and Clarke, M.F. 2003. Prospective identification of tumorigenic breast cancer cells. Proc. Natl. Acad. Sci. U.S.A. 100:39833988. Dalerba, P., Dylla, S.J., Park, I.K., Liu, R., Wang, X., Cho, R.W., Hoey, T., Gurney, A., Huang,

E.H., Simeone, D.M., Shelton, A.A., Parmiani, G., Castelli, C., and Clark, M.F. 2007. Phenotypic characterization of human colorectal cancer stem cells. Proc. Natl. Acad. Sci. U.S.A. 104:10158-10163. O’Brien, C.A., Pollett, A., Gallinger, S., and Dick, J.E. 2007. A human colon cancer cell capable of initiating growth in immunodeficient mice. Nature 445:106-110. Porter, E.H. and Berry, R.J. 1964. The efficient design of transplantable tumour assays. Br. J. Cancer 17:583-595. Ricci-Vitiani, L., Lombardi, D.G., Pilozzi, E., Biffoni, M., Todaro, M., Peschle, C., and DeMaria, R. 2007. Identification and expression of human colon-cancer-initiating cells. Nature 445:111-115. Robinson, J.P., Darzynkiewicz, Z., Dean, P.N., Dressler, L.G., Rabinovitch, P.S., Stewart, C.C., Tanke, H.J., and Wheeless, L.L. (eds.) 2008. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Singh, S.K., Hawkins, C., Clarke, I.D., Squire, J.A., Bayani, J., Hide, T., Henkelman, R.M., Cusimano, M.D., and Dirks, P.B. 2004. Identification of human brain tumor initiating cells. Nature 432:396-401. Todaro, M., Alea, M.P., Di Stefano, A.B., Cammareri, P., Vermeulen, L., Iovino, F., Tripodo, C., Russo, A., Gulotta, G., Medema, J.P., and Stassi, G. 2007. Colon cancer stem cells dictate tumor growth and resist cell death by production of interleukin-4. Cell Stem Cell 1:389-402.


3.1.12 Supplement 7


In Vivo Evaluation of Leukemic Stem Cells through the Xenotransplantation Model

UNIT 3.2

Dominique Bonnet1 1

Cancer Research UK, London Research Institute, Haematopoietic Stem Cell Laboratory, London, United Kingdom

ABSTRACT The xenotransplantation model has been instrumental for the identification and characterization of human leukemic stem cells. This unit describes our current method for the engraftment of human leukemic patients’ samples in the xenotransplanted mouse model. We concentrate uniquely on the model of acute myeloid leukemia, as it was the first type of leukemia for which the xenotransplantation model was developed. Nevertheless, the Basic Protocol could be applied to other sorts of blood disorders. Curr. Protoc. Stem Cell C 2008 by John Wiley & Sons, Inc. Biol. 7:3.2.1-3.2.11. Keywords: hematopoietic stem cell (HSC) r xenotransplantation r immunodeficient mice r leukemic stem cell (LSC)

INTRODUCTION The adaptation of xenotransplantation assays to examine the propagation of acute myeloid leukemia (AML) in vivo has been fundamental in the identification and characterization of leukemia-initiating cells (Lapidot et al., 1994; Bonnet and Dick, 1997). Transplantation of primary AML cells into NOD/SCID mice led to the finding that only rare cells, termed AML-initiating cells (AML-IC), also known as leukemic stem cells (LSC), are capable of initiating and sustaining growth of the leukemic clone in vivo, and serial transplantation experiments showed that AML-IC possess high self-renewal capacity, and thus can be considered to be the leukemic stem cells. The Basic Protocol below describes the most common and simple method to test for the presence of leukemia-initiating cells; this method can also be used to characterize blood disorders. Support protocols describe methods for further purification of leukemic stem cells and the intra-bone injection procedure. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. NOTE: All of the following procedures using human samples should be performed in a level 2 safety tissue culture unit using sterile and proper aseptic techniques.

IDENTIFICATION OF LEUKEMIA STEM CELLS THROUGH XENOTRANSPLANTATION

BASIC PROTOCOL

Xenotransplantation of human cells obtained from AML patients allows identification and characterization of leukemic stem cells.

Materials Immunodeficient mice: NOD/SCID, NOD/SCID-β2 microglobulin null (β2m−/− ), or NOD/SCID IL2R gammanull (Jackson Laboratory) Acidified water: a solution of HCl at a final pH 2.8 to 3.2 Current Protocols in Stem Cell Biology 3.2.1-3.2.11 Published online December 2008 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0302s7 C 2008 John Wiley & Sons, Inc. Copyright

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AML sample: peripheral blood or bone marrow Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) Ammonium chloride solution (Stem Cell Technologies) Fetal bovine serum (FBS; Stem Cell Technologies, cat. no. 06471) Antibodies against human CD45, CD34, CD38, CD33, and CD19 (BD Biosciences Pharmingen) 100 ng/ml 4 ,6-diamidino-2-phenylindole (DAPI; UV excited, Sigma-Aldrich) or TOPRO-3 [HeNe (633-nm) excitable, Molecular Probes] Irradiator: Cesium source is recommended, but an X-ray system or Cobalt source can also be used 29-G, 1/2 -in. needle and insulin syringe (Tyco Healthcare) Dissection tools: scissors and forceps 5-ml snap-top polystyrene tubes Benchtop centrifuge equipped with swing-out bucket rotor for 15- and 50-ml conical tubes Hemacytometer Fluorescent-activated cell sorter, e.g., FACSAria (BD Biosciences) and/or a Moflow (Dako) equipped with 488-nm, 633-nm, and 404-nm lasers 440/40 bandpass (bp) filter for analysis of DAPI, a 530/30 bp filter for FITC, a 575/26 bp for PE, a 695/40 bp for PerCP, and a 660/20 bp for TOPRO-3 Mouse depletion kit (e.g., StemCell Technologies, cat. no. 13066) Additional reagents and equipment for assessing for AML engraftment (Support Protocol 3), parenteral injections (Donovan and Brown, 2006a), euthanasia of mice (Donovan and Brown, 2006b), and performing a cell count using a hemacytometer (UNIT 1C.3) Prepare the immunodeficient mice 1. Keep the animals in a pathogen-free environment. All the NOD/SCID animals used have some impairment of their immune system and may succumb to infections not affecting normal mice. They thus should be kept in pathogen-free status within barrier systems to protect them from current infections.

2. Treat the mice for at least 8 days with acidified water before irradiation. 3. Set the sublethal irradiation dose (between 300 and 375 cGys) on the irradiator used. Success indeed depends on the dose rate/minute of irradiation. If the dose rate/minute is initially too high, try to reduce it by using appropriate shielding. This will reduce the damage to internal organs.

4. Sublethally irradiate the mice before the adoptive transfer of cells. For best results, perform the irradiation 24 hr before the adoptive transfer of cells. The dose of irradiation depends on the mouse strains used and also on the source and irradiator used. It usually varies from 300 to 375 cG (see notes above).

5. Maintain any mouse receiving irradiation on acidified water for 2 weeks following the irradiation dose to prevent diarrhea or weight loss possibly arising due to epithelial damage of the intestines. Any animal showing persistent weight loss >20% of body weight and/or other signs of illness (e.g., rough fur, loss of appetite, inability to groom, and immobility) should be sacrificed. Experience shows that with these measures, the above side effects rarely arise.

Xenotransplantation Model of Leukemia

Prepare AML cells 6. Isolate mononuclear cells (for an example procedure, see Bonnet and Dick, 1997) from fresh peripheral blood or bone marrow samples. Frozen samples can also be used (see Support Protocol).

3.2.2 Supplement 7


7. For prescreening, inject 5 to 10 × 106 mononuclear cells per mouse (4 to 5 mice are tested). Not all AML samples at diagnosis engraft. Thus, for all new samples that arrive in the laboratory, we test the capacity to engraft first by injecting 5 to 10 × 106 viable cells/mouse using intravenous (tail vein) injections. Injection of the cells is done using under 100 to 200 μl/mouse. Usually, cells are resuspended in PBS/2% FBS.

8. After 8 to 12 weeks, sacrifice the mice by cervical dislocation (Donovan and Brown, 2006b) or terminal anesthesia and assess for AML engraftment (see Support Protocol 3). If the AML sample engrafts, purification of the AML-initiating cells (AML-IC) can be performed using Lin, CD34, and CD38 expression and cell sorting (see Support Protocol 3).

Transfer the cells adoptively 9. Subject mice to sublethal irradiation prior to injection of cell preparations (unpurified or purified cell fraction, genetically modified or not). This may be performed on the same day as, or up to 3 days after irradiation.

10. Inject between 106 and 107 cells intravenously via the tail vein (Donovan and Brown, 2006a; maximum volume 1% body weight) using a syringe with a 29-G, 1/2 -in. needle. In some cases, an intra-bone marrow injection (see Support Protocol) might be preferred, especially if a decrease in homing efficiency of the cell transferred is suspected.

Analyze the engraftment 11. Sacrifice mice between 4 to 14 weeks after transplantation using either cervical dislocation (Donovan and Brown, 2006b) or terminal anesthesia. When work under terminal anesthesia is involved, the level of anesthesia should be maintained at sufficient depth for the animal to feel no pain. Blood sampling is not informative as blood samples do not usually match the level of engraftment present in the bone marrow. Indeed, usually few AML cells circulate in the periphery, except in some AML samples (usually samples from patients with poor prognosis) where the AML infiltrates solid organs like spleen and liver. In these cases, the animals become sick and will need to be sacrificed potentially before 10 to 12 weeks.

12. Dissect the femurs, tibias, and iliac crests from the mice and store at room temperature in CMF-PBS before flushing. Remove all connective tissue around the bone.

Prepare bone marrow 13. Place 1 ml of room temperature CMF-PBS in a 5-ml snap-top polystyrene tube. This will be used to flush the bone marrow (see step 15, below).

14. Cut both ends of each bone to provide openings. 15. To flush, insert the 1-ml CMF-PBS-containing insulin syringe into one end of each bone and wash the lumen of the bone with medium pressure. Repeat twice for both ends of the bone or until the bone appears white.

Prepare the cells for FACS analysis 16. To lyse red blood cells, first cool the cell suspension 5 min on ice. Following the cooling of the suspension, add 3 ml of cold ammonium chloride solution to the 1-ml CMF-PBS/cell suspension, mix, and leave for 5 min at 4◦ C. 17. Add 0.5 ml FBS and centrifuge cells 5 min at 380 × g, 4◦ C. 18. Remove the supernatant and resuspend the cells in 1 ml of cold CMF-PBS containing 2% (v/v) FBS.

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Figure 3.2.1

(legend at right)

3.2.4 Supplement 7


19. Count the cells using a hemacytometer (UNIT 1C.3). Store on ice until ready for antibody labeling.

Stain the cells 20. Prepare a mix of human-specific FITC-conjugated anti-CD19, PE-conjugated antiCD33, and PerCP-conjugated anti-CD45 (5 μl/sample/antibody for all stains and compensation/isotype controls). Also, prepare FITC, PE, and PerCP single-color compensation control tubes (5-ml snap-top polystyrene tubes as before) and a combined FITC/PE/PerCP matched isotype control tube. 21. Distribute 15 μl of the antibody mix into each tube of a fresh set of 5-ml tubes for antibody labeling. 22. Dispense 40 μl of each cell suspension into the appropriate antibody labeling tube and leave to label for 30 min at 4◦ C. 23. Wash cells in 2 ml PBS/2% FBS and resuspend in 500 μl of PBS/2% FBS supplemented with a cell impermeant DNA dye for live/dead discrimination, either 100 ng/ml 4 ,6-diamidino-2-phenylindole or TOPRO-3.

Perform FACS analysis 24. To analyze this combination of fluorochromes set up the FACS device with a 488nm excitation source and either a UV or HeNe (633-nm) source depending on your choice of live/dead discriminator. 25. For emission collection ensure you have a 440/40 bandpass (bp) filter for analysis of DAPI, a 530/30 bp filter for FITC, a 575/26 bp for PE, a 695/40 bp for PerCP and a 660/20 bp for TOPRO-3 in place. 26. During FACS analysis, set the photomultiplier gains so that the background signal from the combined isotype control gives 1% to 5% positive cells in each collection channel. 27. Set the compensation amount according the detected spectral overlap. 28. To analyze the engraftment, draw four dotplots as in panels A, B, C, and D in Figure 3.2.1 [440/40 nm versus side-scatter (SSC), forward scatter (FSC) versus SSC, 695/40 nm versus SSC, and 530/30 nm versus 575/26 nm]. 29. First, exclude dead cells from the analysis via a region (R1) around the live, unstained cells as in Figure 3.2.1A. 30. Next, display these cells on a FSC versus SSC plot and select the lymphoid and myeloid cells for further analysis but exclude debris via a region (R2) as in Figure 3.2.1B. 31. Display cells that fall into the first two regions on a 695/40-nm versus SSC plot and draw a generous region around the CD45-PerCP positive cells as in Figure 3.2.1C (R3). Figure 3.2.1 (at left) For analysis of the engraftment by FACS, first the dead cells are excluded using DAPI staining and a live cells region (R1) is drawn (A). Next, these cells are displayed on an FSC versus SSC plot and lymphoid and myeloid cells are selected for further analysis, but debris is excluded via region R2 (B). A generous region is drawn (R3) around the CD45+ cells (C). These CD45+ cells are further analyzed for the expression of CD33 and CD19 (D). The number of events that fall within these regions may be used to calculate the percentage of live, debris-free cells (R2) that are human cells. In addition, the scatter characteristics of cells may be confirmed as consistent with myeloid [high FSC and SSC; (E)] and lymphoid [low FSC and SSC; (F)]. In panel F, there are no CD19+ cells. Engraftment is classed as myeloid leukemia if a population of CD45+ /CD33+ cells is present without an accompanying CD45+ CD19+ /CD33− cell population.

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32. Display these CD45+ cells on a CD19-FITC versus CD33-PE (530/30-nm versus 575/26-nm) dotplot and draw a quadrant to define FITC+ /PE− cells and FITC− /PE+ cell subsets as in Figure 3.2.1D. 33. Use the number of events that fall within these regions to calculate the percentage of live, debris-free cells (R2) that are human cells. In addition, evaluate the scatter characteristics of cells to confirm them as consistent with myeloid (high FSC and SSC, example in Fig. 3.2.1E) and lymphoid (low FSC and SSC, example in Fig. 3.2.1F). In this example, there are no CD19+ cells. Engraftment is classed as myeloid leukemia if a population of CD45+ /CD33+ cells is present without an accompanying CD45+ CD19+ /CD33− cell population. To confirm the leukemic origin of the myeloid cells present in the bone marrow of engrafted mice and if the original AML sample has a known translocation, it is possible to sort human CD45+ cells and perform fluorescent in situ hybridization analysis of the cells or any PCR analysis in search of a fusion product or a mutated gene (NPM, Flt3L, c-kit, WT1, CEBPα). The description of the procedures for FISH or PCR analysis is beyond the scope of this unit.

Perform serial transplantation 34. To test for self-renewal capacity, perform secondary transplantation. Sort human CD45+ cells from either the first primary recipients or enrich them using a mouse depletion kit following the manufacturer’s instructions. 35. Inject the recovered cells into a second recipient using the same protocol as described before (see steps 9 and 10). 36. After 6 to 12 weeks sacrifice the mice and analyze for human cells engraftment the same way as for primary transplantation (see steps 12 to 33). SUPPORT PROTOCOL 1

INTRA-BONE MARROW INJECTION To exclude stem cell homing interference and focus on the intrinsic capacity of a cell to self-renew, a few groups recently developed a highly sensitive strategy based on direct intra-bone marrow (IBM) injection of the candidate human stem cell (Mazurier et al., 2003; Wang et al., 2003; Yahata et al., 2003). IBM injection was found to be a more sensitive and adequate means to measure human HSC capacity. The intra-bone injection technique is performed under a short general anesthesia following a method originally described by Verlinden et al. (1998).

Additional Materials (see Basic Protocol) Anesthetic solution (see recipe) Post-operative analgesic (Vetergesic; Alstoe Animal health), diluted 1/10 in PBS and injected at 100 μl subcutaneously per mouse 29-G, 1/2 -in. needle (or 25-G needle) and insulin syringe (Tyco Healthcare) Anesthetize the mice 1. Inject the mice intraperitoneally with a dose of 0.2 to 0.25 ml of anesthetic solution. General anesthesia suppresses the heat-regulating mechanisms of the body. This is overcome by intra- and post-operative maintenance of body temperature in appropriate thermostatically controlled incubators or by other heat sources.


2. Insert a syringe with a 25-G needle (maximum) into the joint surface of the right or left tibia/femur, and inject up to 40 μl cell suspension into the bone marrow cavity of the tibia or femur.

3.2.6 Supplement 7


3. During recovery, keep the animals under regular observation until full mobility is regained. 4. At this stage, provide at least one dose (100 μl) of post-operative analgesic (Vetergesic diluted 1/10) subcutaneously following bone marrow injection.

THAWING AML CELLS While freshly obtained AML cells are desirable for xenotransplantation, thawed frozen cells are also suitable, and it may be more convenient to collect and freeze the cells until one is ready for the transplantation experiment.

SUPPORT PROTOCOL 2

Materials AML cells, frozen in 1.8- to 2-ml cryovials DNase (Sigma, cat. no. D4513), thawed Fetal bovine serum (FBS; Stem Cell Technologies, cat. no. 06471) or any other suppliers Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) 37◦ C water bath 50-ml centrifuge tubes, sterile Table-top centrifuge equipped with swing-out bucket rotor for 15- and 50-ml conical tubes Cell strainer Additional reagents and equipment for counting cells (UNIT 1C.3) NOTE: Before thawing the samples, ensure the water bath is at 37◦ C. NOTE: Before starting, ensure that the DNase is completely thawed. 1. Rapidly thaw the AML cells (cryovial of 1.8- to 2-ml) in the 37◦ C water bath. There are no commercial suppliers of frozen AML. These samples can be obtained from clinics after informed consent from the patients has been obtained.

2. Add 100 ml DNase (1 mg/ml) dropwise into the cryovial. 3. Mix gently, wait 1 min, and transfer cells into a sterile 50-ml centrifuge tube. 4. Gently add 1 ml heat-inactivated, pure FBS dropwise, mix gently, and wait 1 min. 5. Slowly add 10 ml CMF-PBS/2% FBS and wait 1 min. 6. Slowly add up to 30 ml CMF-PBS/2% FBS to fill the tube. 7. Centrifuge 5 min at 200 × g, 4◦ C. 8. Resuspend in 1 ml CMF-PBS/2% FBS. 9. Filter using a cell strainer if needed (if cells are clumping). 10. Count viable cells (UNIT 1C.3) and use for purification or adoptive transfer protocols.

PURIFICATION STRATEGY Mononuclear cells from AML samples can be stained using a combination of antibodies. The most commonly used are the lineage cocktail antibodies (BD, cat. no. 340546), antiCD34 and anti-CD38. Stain the cells 25 to 30 min at 4◦ C in the presence of 5 μl/million of each of these antibodies. After the incubation period, centrifuge the cells 5 min at 380 × g, 4◦ C. Discard the supernatant, resuspend the cells in 1 ml CMF-PBS/2% FBS. Repeat this wash procedure one more time. After washing, the cells are ready to be sorted through a cell sorter (FACS Aria, BD, or equivalent).

SUPPORT PROTOCOL 3

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Anesthetic solution Mix 1 ml of Ketase solution (Fort Dodge Animal Health) with 0.5 ml of 2% Rompun solution (Bayer plc) and dilute with 8.5 ml of CMF-PBS. Store up to 1 month at 4◦ C. Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) Prepare 10× stock with 1.37 M NaCl, 27 mM KCl, 100 mM Na2 HPO2 , and 18 mM KH2 PO4 (adjust to pH 7.4 using HCl, if necessary) and autoclave. Store up to 2 months at room temperature. Prepare working solution by dilution of one part with nine parts water.



AML-IC can be prospectively identified and purified as CD34+ /CD38− cells in AML patient samples, regardless of the phenotype of the bulk blast population, and these cells represent the only AML cells capable of self-renewal (Bonnet and Dick, 1997). The phenotype of AML-IC has been extended to include the expression of CD123 but the absence of CD71, HLA-DR, and CD117 (Blair et al., 1997, 1998; Blair and Sutherland, 2000; Jordan et al., 2000). In a recent study, this phenotype was further extended to include expression of CD33 and CD13 on AML-IC for the vast majority of patients (Taussig et al., 2005). Hence, the immunophenotype of the leukemic stem cell as defined by in vivo propagation is CD34+ /CD38− /CD71− / HLA-DR− /CD117− /CD33+ /CD13+ /CD123+ . However, the exclusivity of some of these markers is debatable. For instance, CD123 is indeed expressed on AML-IC, but it is also expressed on the vast majority of AML blasts (D. Bonnet, unpub. observ.) from most patients, and hence could be excluded from the above phenotype of AML-IC. Considerable heterogeneity within the AML-IC compartment exists. Lentiviral gene marking to track the behavior of individual LSCs, following serial transplantation, has revealed heterogeneity in their ability to selfrenew, similar to what is seen in the normal HSC compartment (Hope et al., 2004). Furthermore, using the NOD/SCID IL2Rgnull mice (see Anticipated Results) pretreated with anti-CD122, we show that in some patients the LSC activity can be detected in a CD34+ CD38+ population (Taussig et al., 2008). Thus, there is not a universal protocol

to purify for LSC in AML. Each AML patient sample should be tested first for its ability to engraft (prescreening) and secondly for identifying the nature of the cells responsible for repopulating ability. Secondary transplantation should also be used to test for the self-renewal ability of the LSC.

Critical Parameters By LSC, we refer to a cell that has selfrenewal and differentiation potential and is able to reinitiate the leukemia when transplanted into NOD/SCID mice. This definition does not preclude the nature of the cells being transformed. The confusion regarding the origin of the AML-IC may be due to the extreme heterogeneity of AML. Given the various possible routes to AML from a normal hematopoietic cell, it is not surprising that there is great heterogeneity in AML. Indeed, AML may be thought of as a large collection of different diseases that merely share similar characteristics. Indeed, the most effective risk stratification approach so far has been to examine the genetic abnormalities associated with a particular case of AML and compare it to previous experience with AML cases with the same abnormality (Grimwade et al., 1998, 2001). Although cytogenetic analysis allows the definition of the hierarchical groups with favorable, intermediate, and poor prognosis, the intermediate risk group contains patients with variable outcomes. Assessing the prognosis of this large group of patients is currently difficult.

Troubleshooting It is usually straightforward to detect AML engraftment. The human cells present express

3.2.8 Supplement 7


human CD45+ and the pan-myeloid marker CD33 without detection of lymphoid markers (CD19). Nevertheless, it happens that in some cases a chimeric engraftment can be detected, indicating that both human normal and leukemic cells are present in the engrafted mouse. In this case, it is usually important to confirm the leukemic origin of the CD33 subfraction by performing either FISH analysis or PCR for the mutations present in the original patient samples. Human CD45+ can be sorted based on CD19 and CD33, and both fractions should be tested for the presence of leukemic cells. Thus, it is not sufficient when screening for human AML engraftment to only test for the presence of human CD45+ cells, as in some cases these human cells could be exclusively normal cells.

Anticipated Results From AML patients’ samples at diagnosis, the capacity to engraft in the xenotransplantation model is usually ∼65% to 70%. Thus, there are still 25% to 30% of patients for which no engraftment could be detected after 10 to 12 weeks. The ability of a particular AML to engraft in the xenotransplantation model is related to the prognosis of individual AML cases (Pearce et al., 2006). Specifically, examination of the follow-up results from younger patients with intermediate-risk AML revealed a significant difference in overall survival between NOD/SCID-engrafting and non-engrafting cases. No differences have been detected between engrafting and nonengrafting cases in various engraftment variables including: homing ability, AML-IC frequency, and immune rejection by the host or alternative tissue sources. Hence, the ability to engraft NOD/SCID recipients seems to be an inherent property of the cells that is directly related to prognosis. Other mouse models have been developed to support the growth of human hematopoietic cells but less is known about the ability of these new models to sustain AML engraftment. The NOD/SCID-β2 microglobulin null (β2m−/− ) mouse has an additional defect in NK cell activity and is more tolerant of human grafts than the NOD/SCID model (Christianson et al., 1997; Kollet et al., 2000; Glimm et al., 2001). However, the percentage of AML samples that engraft in β2m−/− is similar to the level achieved using the NOD/SCID mice. Thus, it does not appear that the β2m−/− is superior for the engraftment of AML samples (Pearce et al., 2006). Furthermore, both NOD/SCID and β2m−/− are susceptible to developing lymphomas over time,

limiting their lifespan and preventing longterm reconstitution assessment. These hurdles have recently been overcome in three new strains: NOD/Shi-Scid IL2Rgnull (Yahata et al., 2002; Hiramatsu et al., 2003), NOD/SCID IL2Rgnull (Ishikawa et al., 2005; Shultz et al., 2005), and BALB/c-Rag2null IL2Rgnull (Traggiai et al., 2004), which all lack the IL-2 family common cytokine receptor gamma chain gene. The absence of functional receptors for IL2, IL-7, and other cytokines may prevent the expansion of NK cells and early lymphoma cells in NOD/SCID IL2Rgnull mice, resulting in better engraftment of transplanted human cells and longer lifespan of the mice. It was reported recently that human HSCs and progenitor cells engraft successfully in these mice and produce all human myeloid and lymphoid lineages. T and B cells migrate into lymphoid organs and mount HLA-dependent allogeneic responses, and generate antibodies against T cell–dependent antigens such as ovalbumin and tetanus toxin (Traggiai et al., 2004; Ishikawa et al., 2005). However, preliminary testing in our group seems to indicate that the NOD/SCID IL2Rgnull mice are not superior for leukemic engraftment to NOD/SCID or the NOD/SCID-β2m−/− mice. Thus, intrinsic properties of AML cells will dictate whether or not the cells will engraft.

Time Considerations It usually takes 4 to 6 weeks to detect engraftment. Nevertheless, we have seen in the past that the kinetics of AML engraftment, in contrast to normal stem cells, is usually slower, and thus it is better to wait 10 to 12 weeks for estimating the engraftment of AML samples. It sometimes happens, especially with poor cytogenetic samples that the mice get sick after only 3 to 4 weeks due to an infiltration of AML cells into solid organs (like spleen, liver, kidney). In this case, the mice have to be sacrificed earlier.

Acknowledgments The author would like to thank Dr. Daniel Pearce for his assistance in the preparation of this manuscript.

Literature Cited Blair, A., Hogge, D.E., Ailles, L.E., Lansdorp, P.M., and Sutherland, H.J. 1997. Lack of expression of Thy-1 (CD90) on acute myeloid leukemia cells with long-term proliferative ability in vitro and in vivo. Blood 89:3104-3112. Blair, A., Hogge, D.E., and Sutherland, H.J. 1998. Most acute myeloid leukemia progenitor cells with long-term proliferative ability in vitro and

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in vivo have the phenotype CD34(+)/CD71 (-)/HLA-DR. Blood 92:4325-4335. Blair, A. and Sutherland, H.J. 2000. Primitive acute myeloid leukemia cells with long-term proliferative ability in vitro and in vivo lack surface expression of c-kit (CD117). Exp. Hematol. 28:660-671. Bonnet, D. and Dick, J.E. 1997. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat. Med. 3:730-737. Christianson, S.W., Greiner, D.L., Hesselton, R.A., Leif, J.H., Wagar, E.J., Schweitzer, I.B., Raian, T.V., Gott, B., Roopenian, D.C., and Shultz, L.D. 1997. Enhanced human CD4+ T cell engraftment in beta2-microglobulin-deficient NODscid mice. J. Immunol. 158:3578-3586. Donovan, J. and Brown, P. 2006a. Parenteral injections. Curr. Protoc. Immunol. 73:1.6.1-1.6.10. Donovan, J. and Brown, P. 2006b. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Glimm, H., Eisterer, W., Lee, K., Cashman, J., Holyoake, T. L., Nicolini, F., Shultz, L.D., von Kalle, C., and Eaves, C.J. 2001. Previously undetected human hematopoietic cell populations with short-term repopulating activity selectively engraft NOD/SCID-beta2 microglobulinnull mice. J. Clin. Invest. 107:199-206. Grimwade, D., Walker, H., Oliver, F., Wheatley, K., Harrison, C., Harrison, G., Rees, J., Hahn, I., Stevens, R., Burnett, A., and Gladstone, A. 1998. The importance of diagnostic cytogenetics on outcome in AML: Analysis of 1,612 patients entered into the MRC AML 10 trial. The Medical Research Council Adult and Children’s Leukaemia Working Parties. Blood 92:23222333. Grimwade, D., Walker, H., Harrison, G., Oliver, F., Chatters, S., Harrison, C. J., Wheatley, K., Burnett, A.K., and Goldstone, A.H.; Medical Research Council Adult Leukemia Working Party. 2001. The predictive value of hierarchical cytogenetic classification in older adults with acute myeloid leukemia (AML): Analysis of 1065 patients entered into the United Kingdom Medical Research Council AML11 trial. Blood 98:1312-1320. Hiramatsu, H., Nishikomori, R., Heike, T., Ito, M., Kobayashi, K., Katamura, K., and Nakahata, T. 2003. Complete reconstitution of human lymphocytes from cord blood CD34+ cells using the NOD/SCID/gammacnull mice model. Blood 102:873-880. Hope, K. J., Jin, L., and Dick, J. E. 2004. Acute myeloid leukemia originates from a hierarchy of leukemic stem cell classes that differ in self-renewal capacity. Nat. Immunol. 5:738743.


Ishikawa, F., Yasukawa, M., Lyons, B., Yoshida, S., Miyamoto, T., Yoshimoto, G., Watanabe, T., Akashi, K., Shultz, L.D., and Harada, M. 2005. Development of functional human blood and immune systems in NOD/SCID/IL2 receptor {gamma} chain(null) mice. Blood 106:15651573.

Jordan, C.T., Upchurch, D., Szilvassy, S.J., Guzman, M.L., Howard, D.S., Pettigrew, A.L., Meyerrose, T., Rossi, R., Grimes, B., Rizzeri, D.A., Luger, S.M., and Phillips, G.L. 2000. The interleukin-3 receptor alpha chain is a unique marker for human acute myelogenous leukemia stem cells. Leukemia 14:1777-1784. Lapidot, T., Sirard, C., Vormoor, J., Murdoch, B., Hoang, T., Caceres-Cortes, J., Minden, M., Paterson, B., Caligiuri, M.A., and Dick, J.E. 1994. A cell initiating human acute myeloid leukaemia after transplantation into SCID mice. Nature 367:645-648. Kollet, O., Peled, A., Byk, T., Ben-Hur, H., Greiner, D., Shultz, L., and Lapidot, T. 2000. beta2 microglobulin-deficient (B2m(null)) NOD/SCID mice are excellent recipients for studying human stem cell function. Blood 95:3102-3105. Mazurier, F., Doedens, M., Gan, O.I., and Dick, J.E. 2003. Rapid myeloerythroid repopulation after intrafemoral transplantation of NOD-SCID mice reveals a new class of human stem cells. Nat. Med. 9:959-963. Pearce, D.J., Taussig, D., Zibara, K., Smith, L.L., Ridler, C.M., Preudhomme, C., Young, B.D., Rohatneer, A.Z., Lister, T.A., and Bonnet, D. 2006. AML engraftment in the NOD/SCID assay reflects the outcome of AML: Implications for our understanding of the heterogeneity of AML. Blood 107:1166-1173. Shultz, L.D., Lyons, B.L., Burzenski, L.M., Gott, B., Chen, X., Chaleff, S., Kotb, M., Gillies, S.D., King, M., Mangada, J., Greiner, D.L., and Handgretinger, R. 2005. Human lymphoid and myeloid cell development in NOD/LtSz-scid IL2R gamma null mice engrafted with mobilized human hematopoietic stem cells. J. Immunol. 174:6477-6489. Taussig, D.C., Pearce, D.J., Simpson, C., Rohatiner, A.Z., Lister, T.A., Kelly, G., Luongo, J.L., Danet-Desnoyers, G.A., and Bonnet, D. 2005. Hematopoietic stem cells express multiple myeloid markers: Implications for the origin and targeted therapy of acute myeloid leukemia. Blood 106:4086-4092. Taussig, D.C., Miraki-Moud, F., Anjos-Afonso, F., Pearce, D.J., Allen, K., Ridler, C., Lillington, D., Oakervee, H., Cavenagh, J., Agrawal, S.G., Lister, T.A., Gribben, J.G., and Bonnet, D. 2008. Anti-CD38 antibody-mediated clearance of human repopulating cells masks the heterogeneity of leukemia-initiating cells. Blood 112:568575. Traggiai, E., Chicha, L., Mazzucchelli, L., Bronz, L., Piffaretti, J.C., Lanzavecchia, A., and Manz, M.G. 2004. Development of a human adaptive immune system in cord blood cell-transplanted mice. Science 304:104-107. Verlinden, S.F., van Es, H.H., and van Bekkum, D.W. 1998. Serial bone marrow sampling for long-term follow up of human hematopoiesis in NOD/SCID mice. Exp. Hematol. 26:627-630. Wang, J., Kimura, T., Asada, R., Harada, S., Yokota, S., Kawamoto, Y., Fijimura, Y., Tsuji,

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T., Ikehara, S., and Sonoda, Y. 2003. SCIDrepopulating cell activity of human cord bloodderived CD34- cells assured by intra-bone marrow injection. Blood 101:2924-2931. Yahata, T., Ando, K., Nakamura, Y., Ueyama, Y., Shimamura, K., Tamaoki, N., Kato, S., and Hotta, T. 2002. Functional human T lymphocyte development from cord blood CD34+ cells in nonobese diabetic/Shi-scid, IL-2 receptor gamma null mice. J. Immunol. 169:204-209. Yahata, T., Ando, K., Sato, T., Miyatake, H., Nakamura, Y., Muguruma, Y., Kato, S., and Hotta, T. 2003. A highly sensitive strategy for SCID-repopulating cell assay by direct injection of primitive human hematopoietic cells into NOD/SCID mice bone marrow. Blood 101:2905-2913.

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Culture and Isolation of Brain Tumor Initiating Cells

UNIT 3.3

Monika Lenkiewicz,1 Na Li,1 and Sheila K. Singh1 1

McMaster University, Hamilton, Ontario, Canada

ABSTRACT This unit describes protocols for the culture and isolation of brain tumor initiating cells (BTIC). The cancer stem cell (CSC) hypothesis suggests that tumors are maintained exclusively by a rare fraction of cells that have stem cell properties. We applied culture conditions and assays originally used for normal neural stem cells (NSCs) in vitro to a variety of brain tumors. The BTIC were isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133. Only the CD133+ brain tumor fraction contains cells capable of sphere formation and sustained self-renewal in vitro, and tumor initiation in NOD-SCID mouse brains. Therefore, CD133+ BTICs satisfy the definition of cancer stem cells in that they are able to generate a replica of the patient’s tumor and they exhibit self-renewal ability through serial retransplantation. This established that only a rare subset of brain tumor cells with stem cell properties are tumor-initiating, and, in this unit, we describe their culture and isolation. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 11:3.3.1-3.3.10. Keywords: brain tumor initiating cells (BTICs) r tumor sphere culture r CD133 r cell sorting r cancer stem cell (CSC)

INTRODUCTION In this unit, protocols for the culture and isolation of brain tumor initiating cells (BTICs) are described. The cancer stem cell (CSC) hypothesis suggests that tumors are maintained exclusively by a rare fraction of cells that have stem cell properties. Here, we discuss the methods that we first used to prospectively identify and enrich for a subpopulation of human BTICs that exhibit the stem cell properties of proliferation, self-renewal, and differentiation in vitro (Singh et al., 2003) and in vivo (Singh et al., 2004). We applied culture conditions and assays originally used to characterize normal neural stem cells (NSCs) in vitro (Reynolds and Weiss, 1992; Tropepe et al., 1999) to a variety of pediatric and adult brain tumors. The BTIC were exclusively isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133 (Yin et al., 1997; Yu et al., 2002). Only the CD133+ brain tumor fraction contains cells that are capable of sphere formation and sustained self-renewal in vitro, as well as tumor initiation in NOD-SCID mouse brains. Therefore, CD133+ BTICs satisfy the definition of cancer stem cells in that they are able to generate a replica of the patient’s tumor and they exhibit self-renewal ability both in vitro and in vivo through serial retransplantation (Bonnet and Dick, 1997; Reya et al., 2001). This formally established that only a rare subset of brain tumor cells with stem cell properties are tumor-initiating. This unit begins with a method for the culture of tumor spheres from human brain tumors (Basic Protocol 1) and follows with a protocol for the prospective isolation of BTICs from these cultures by magnetic bead sorting (Basic Protocol 2) or fluorescence activated sorting (Alternate Protocol) for CD133. NOTE: The following procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. Current Protocols in Stem Cell Biology 3.3.1-3.3.10 Published online October 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc0303s11 C 2009 John Wiley & Sons, Inc. Copyright

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3.3.1 Supplement 11

NOTE: All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. BASIC PROTOCOL 1

CULTURE OF TUMOR SPHERES FROM HUMAN BRAIN TUMORS This protocol is adapted from those previously established for isolation of neural stem cells as neurospheres (Reynolds and Weiss, 1992, 1996; Reynolds et al., 1992), and has been applied to the culture of human brain tumors. We use this culture method specifically to select for cell populations within human brain tumors that possess stem cell properties. Serum-free medium (SFM) allows for the maintenance of an undifferentiated stem cell state, and the addition of bFGF and EGF allows for the proliferation of multipotent, selfrenewing, and expandable tumor spheres. The medium on these tumor sphere cultures should be changed every other day, and when primary tumor spheres reach a critical size of >100 μm, they may be passaged to secondary spheres based on the principles of the neurosphere assay. The frequency of the stem cell population within the brain tumor can be determined by primary sphere formation assay, and the minimal frequency of repopulating tumor sphere cells within the culture can be estimated by serial sphere formation through limiting dilution analysis (Tropepe et al., 1999).

Materials Hi/low aCSF (see recipe) 95% O2 /5% CO2 Enzyme digestive mix for tumors (see recipe) or high-performance liquid chromatography–purified collagenase/dispase cocktail (e.g., Liberase Blendzyme 3 from Roche) Human brain tumor specimen Tumor sphere medium (see recipe) Soybean trypsin inhibitor (from Glycine max; Sigma, cat. no. T9003) 10 ng/μl leukemia inhibitor factor (LIF) stock solution (see recipe) 10 ng/μl recombinant human basic fibroblast (bFGF) stock solution (see recipe) 10 ng/μl recombinant human epidermal growth factor (EGF) stock solution (see recipe) 0.22-μm pore size, 150-ml filter system (Corning) Oxygen tank setup in laminar flow hood to allow for sterile oxygenation of solutions 15- and 50-ml conical centrifuge tubes 0.22-μm syringe filter units (Millipore) 10-cm2 tissue culture–grade plates (Falcon) or 100-mm Ultra Low Attachment Culture dishes (Corning) Fine sterile scissors and forceps Incubator-shaker (VWR Scientific) Tabletop centrifuge 70-μm cell strainer (BD Falcon) Dissociate human brain tumor tissue 1. Prepare 125 ml of hi/low aCSF as described in Reagents and Solutions, and filter sterilize using a 0.2-μm filter and 150-ml filter system. Culture and Isolation of Brain Tumor Initiating Cells

2. Bubble aCSF with sterile 95% O2 /5% CO2 for 15 min and place in 37◦ C water bath.

3.3.2 Supplement 11


3. Freshly prepare the enzyme digestive mix for tumors (see Reagents and Solutions) by weighing out the enzymes (and kynurenic acid) into separate 15-ml centrifuge tubes. Resuspend each enzyme/reagent in 10-ml of hi/low aCSF, vortex thoroughly, and filter all of the components through a 0.22-μm syringe filter into one sterile 50-ml centrifuge tube. Alternatively, measure high performance liquid chromatography– purified collagenase/dispase cocktail (Liberase Blendzyme 3) according to the manufacturer’s instructions, and resuspend in aCSF. If using enzyme digestive mix for tumors, final working concentrations are: trypsin 1.33 mg/ml, hyaluronidase 0.67 mg/ml, and kynurenic acid 0.1 to 0.17 mg/ml. If using the alternate method with Liberase Blendzyme 3, the final working concentration is 0.2 Wünsch units/ml in a total of 15 ml aCSF.

4. In sterile hood, wash tumor specimen in a 10-cm2 plate filled with hi/low aCSF, transferring sequentially to new 10-cm2 plates filled with aCSF until excess blood is thoroughly washed out. The authors have tested 12 brain tumor subtypes (Singh et al., 2003, 2004) and they all had some proportion of BTICs; sample size can be very small and BTICs are still isolated, so there is no minimum sample size. BTIC yield directly correlates with grade of tumor, i.e., there is lower BTIC yield with benign low-grade tumors and higher yield with biologically aggressive malignant tumors (Singh et al., 2003, 2004).

5. Using sterile fine scissors and forceps, cut tumor into 1-mm3 pieces in a 10-cm2 tissue culture plate with 2 to 3 ml of enzymatic digestion mixture (either the enzyme/ kynurenic acid mix used in step 3, or Liberase Blendzyme 3). 6. Collect tumor pieces with 10-ml pipet, pipetting up and down and dispensing the tumor fragments into 30 ml of the enzymatic digestion mixture used in step 3 or 15 ml Liberase Blendzyme 3. 7. Digest at 37◦ C for 30 to 90 min (depending on tumor size) with gentle mixing in an incubator-shaker. The incubator-shaker provides superior digestion and subsequent yield of cells compared to a rocker. If using Liberase Blendzyme 3, the digestion period can be reduced to 15 to 30 min.

Stop enzymatic reaction Steps 8 to 10 are skipped if using Liberase Blendzyme 3. In that case, simply filter the cell digest from step 7 through a 70-μm cell strainer and proceed to step 11. 8. Freshly prepare the trypsin inhibitor solution by resuspending 35 mg of trypsin inhibitor in 5 ml tumor sphere medium (without LIF, bFGF, or EGF) and filtering through a 0.2-μm filter. 9. Centrifuge cells 3 min at ∼450 × g, room temperature, and take off as much supernatant as possible without dislodging the tumor tissue at the bottom of the tube. 10. Add the 5 ml of trypsin inhibitor solution from step 8, mix well, and filter through a 70-μm cell strainer.

Plate cells in tumor sphere medium with growth factors 11. Centrifuge cells 3 min at ∼450 × g, room temperature. Aspirate all of the supernatant and resuspend in 10 to 12 ml tumor sphere medium supplemented with 10 ng/ml LIF (add from 10 ng/μl stock), 20 ng/ml bFGF (add from 10 ng/μl stock), and 20 ng/ml EGF (add from 10 ng/μl stock). 12. Plate cells in 10-cm2 dishes at 2 × 105 cells per cm2 , in 10 to 12 ml of tumor sphere medium with LIF, bFGF, and EGF. Incubate in a 37◦ C 5% CO2 incubator.

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A 100x

400x

0.05 mm

0.05 mm

B 100x

200x

0.05 mm

0.05 mm

Figure 3.3.1 Brain tumor initiating cells forming sphere-like structures in vitro. (A) Tumor spheres of anaplastic medulloblastoma. (B) Tumor spheres of metastatic melanoma.

13. Feed cells every other day by centrifuging 3 min at 450 × g, room temperature, aspirating the medium, and replacing it with fresh tumor sphere medium supplemented with fresh LIF, bFGF, and EGF, as described in step 11. 14. When number of spheres per plate has doubled or spheres are consistently >120 μm in size, split cultures by repeating steps 11 to 12. Figure 3.3.1 shows brain tumor initiating cells forming sphere-like structures in vitro.

BASIC PROTOCOL 2


ENRICHMENT OF BTICs BY MAGNETIC BEAD SORTING FOR CD133 This protocol is used for the prospective isolation or enrichment of BTICs from tumor sphere cultures, which constitute primary human brain tumor cells, cultured as per Basic Protocol 1. Cell sorting is performed as soon as tumor spheres begin to form in culture, and is optimally performed within 1 to 24 hr after initial cell culture. Cells must be in single-cell suspension for optimal sorting, and spheres are gently triturated or chemically dissociated prior to cell sorting, by methods detailed below. Cell sorting for CD133 can be performed either by magnetic bead cell sorting (MBCS; this protocol) or by fluorescence activated cell sorting (FACS; Alternate Protocol), and we provide methods for each of these options. Considerations of experimental timing, cell sorter availability at short notice or at night, cell viability after processing through FACS, individual tumor characteristics, overall cell number, and inter-user reliability can help to discern which method of cell sorting should be used for each tumor sphere sample.

3.3.4 Supplement 11


Materials Primary human brain tumor cells growing in culture (Basic Protocol 1) Incubation buffer: phosphate buffered saline (PBS; Invitrogen, cat. no. 14190-144)/0.5% (w/v) BSA with (or without; see below) 2 mM disodium EDTA, pH 7.2 FcR blocking reagent (Miltenyi Biotec) Phosphate-buffered saline (Invitrogen, cat. no. 14190-144) MACS CD133 Cell Isolation Kit (Miltenyi Biotec) consisting of: beads conjugated to monoclonal mouse anti–human CD133 Microbeads, Isotype IgG1 magnetic cell separator (MiniMACS column magnet) MS separation columns CD133-2-PE antibody mouse IgG2b-PE isotype control antibody Tumor sphere medium (see recipe) 4% (w/v) paraformaldehyde (optional) 15-ml conical centrifuge tubes (Falcon or equivalent) Flame-narrowed pipets 70-μm cell strainer 6-well Ultra Low Cluster plates (Corning) Flow cytometry tubes (BD Falcon 352058) Additional reagents and equipment for counting viable cells by trypan blue exclusion (UNIT 1C.3) and flow cytometry (Robinson et al., 2009) NOTE: Work quickly and keep cells/buffer (not culture media) cold (4◦ to 6◦ C).

Prepare cell suspensions 1. Place 2 × 105 cultured tumor spheres in a 15-ml centrifuge tube. Centrifuge cells 3 min at ∼450 × g, room temperature, and remove medium. Resuspend cell pellet in 1 ml of incubation buffer. Do not use EDTA if working with Notch pathway molecules, due to potential interactions. RBC contamination will decrease purity and cause tumor cell death upon RBC lysis; to remove RBCs if specimen is vascular, treat the cells with an RBC lysis buffer (StemCell Technologies, cat. no. 67850).

2. Triturate gently with flame-narrowed pipet or micropipet tip (if not at single-cell suspension already). 3. Filter through 70-μm cell strainer, and count viable cells using a hemacytometer and trypan blue (UNIT 1C.3). Gently triturate and strain cells just before flow to avoid clumping. You may use a flamenarrowed or regular pipet or micropipet; we avoid excessive trituration to minimize trauma to the cells.

4. Aliquot equal amounts of cell suspension into four microcentrifuge tubes as follows:

negative control (unstained) (A1) isotype control (A2) pre-sort CD133-1 staining of specimen (A3) bulk of the cells in the last aliquot, for bead sorting (A4). Perform magnetic separation 5. Resuspend A1 and A4 in 300 μl incubation buffer and add 100 μl FcR blocking reagent to A4. If cell number in A4 is >5 × 106 , divide into two tubes and treat both the same way. Resuspend A2 and A3 in 500 μl PBS. Keep aliquots A1, A2, and A3 on ice. Current Protocols in Stem Cell Biology

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6. Add 1 μl beads per 106 cells to tube A4. Mix beads well in A4 and leave at 4◦ C (in the refrigerator in dark) for 30 min. 7. After 30 min, take the A4 aliquot and place the microcentrifuge tube into a MACS separator magnet. Use LS columns or MS columns provided with the kit, based on cell number. 8. Rinse column with 3 ml incubation buffer for LS column. Add 3 ml cell suspension onto the column, and allow negative cells to pass through. Collect negative cells in a 15-ml centrifuge tube. 9. Wash four times, each time with 3 ml incubation buffer. Pool with the negative cells in the 15-ml centrifuge tube from step 8. 10. Remove column from separator and place on a clean 15-ml centrifuge tube. Flush out positive fraction with 3 ml incubation buffer by firm application of hand to column. 11. Repeat, and apply column one more time to resuspended CD133+ cells, to purify this population further if cell numbers permit. Reapplying the cells to the column increases the purity of the CD133+ cells to ∼95%.

12. Centrifuge the sorted A4 aliquots, CD133− from steps 8 and 9 and CD133+ from step 11, 3 min at 450 × g, room temperature, and remove the supernatant. Wash by adding 2 ml incubation buffer, centrifuging again as before, and removing the supernatant. Resuspend each pellet in 3 ml tumor sphere medium. 13. Plate 2–5 × 104 cells per well in 3 ml tumor sphere medium per well of a 6-well plate. Re-equilibrate immediately to 37◦ C.

Perform a purity check by immunostaining and flow cytometric analysis 14. Take a 10-μl aliquot from the last CD133+ and CD133− aliquots (step 12) and resuspend each in 500 μl PBS. 15. Add 5 μl CD133-2-PE antibody to each sample, and incubate at 4◦ C for 30 min. These specimens will be taken to the flow cytometry lab for a purity check of the CD133+ and CD133− sorted populations.

16. Add 5 to 10 μl of CD133-2-PE antibody to tube A2 and 5 to 10 μl isotype IgG2b-PE control antibody to tube A3 (see step 4). Incubate at room temperature for 15 to 30 min. Also incubate the unstained A1 control and carry it through the remaining steps. 17. Centrifuge aliquots A2 and A3 3 min at ∼450 × g, room temperature, and remove the supernatants. Wash by adding 10 to 20 pellet volumes of PBS, centrifuging again as before, and removing the supernatant. Finally, resuspend in 250 μl PBS. The cells should be washed as thoroughly as possible after staining; however, this must be balanced with the potential for cell loss.

18. Optional: Post-fix A2, A3, and A1 aliquots and the two purity check aliquots from A4 in 2% (w/v) paraformaldehyde (i.e., add 250 μl 4% paraformaldehyde to each aliquot). 19. Triturate each sample gently and transfer to flow cytometry tubes, preferably through strainer caps to remove clumps. Culture and Isolation of Brain Tumor Initiating Cells

20. Analyze the five samples by flow cytometry (Robinson et al., 2009). See Figure 3.3.2 for sample results.

3.3.6 Supplement 11


A tumor cells

negative control (A1) IgG2b isotype control (A2) CD133-2-PE (A3)

magnetic sorting (A4) (Miltenyi CD 133 isolation kit) CD133 cells

CD133 cells

purity check (flow cytometry)

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64 95% ( )

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Figure 3.3.2 (A) Histogram of magnetic bead separation protocol for CD133. (B) Histograms of flow cytometry for the unsorted tumor cells (left) and sorted CD133+ cells (right) from a 49-year-old female metastatic melanoma patient. Corresponding isotype controls were overlaid as the unshaded histograms.

ENRICHMENT OF BTICs BY FLUORESCENCE ACTIVATED CELL SORTING

ALTERNATE PROTOCOL

Cell preparation and staining are performed as per the magnetic separation protocol (Basic Protocol 2), with the exception that the A4 aliquot, containing the bulk of the cells, is used for fluorescence activated cell sorting (FACS; Robinson et al., 2009). This aliquot should be resuspended in 300 μl PBS with 100 μl FcR blocking reagent. If the cell number in A4 is >5 × 106 , divide the suspension into two tubes and treat both the same way. Incubate A4 with a 1:100 dilution of CD133-2-PE antibody and leave at 4◦ C (in refrigerator in dark) for 30 min. Wash and centrifuge in 10 to 20 volumes of PBS as described in Basic Protocol 2, and resuspend A4 cells in 250 μl PBS. Take A4 aliquot to your FACS operator, with aliquots A1 through A3 prepared as in Basic Protocol 2 for controls.


Enzyme digestive mix for tumors 40 mg bovine pancreas trypsin (Sigma, cat no. T9201) 20 mg bovine testis hyaluronidase (Sigma, cat no. H3884) 3 to 5 mg kynurenic acid (Sigma, cat no. K3375) Current Protocols in Stem Cell Biology

Cancer Stem Cells continued

3.3.7 Supplement 11

Resuspend these three components in 30 ml of warm (37◦ C) hi/low aCSF and filter through a 0.2-μm filter. This enzyme digestive mix must be made up fresh prior to use.

Hi/low artificial cerebral spinal fluid (aCSF) For a total of 125 ml aCSF, combine the following: 7.75 ml 2 M NaCl 0.625 ml 1 M KCl 0.4 ml 1 M MgCl2 21.125 ml 155 mM NaHCO3 1.25 ml 1 M glucose 0.1157 ml 108 mM CaCl2 93.73 ml H2 O Hi/low aCSF can be prepared and stored at 4◦ C for up to 6 months (Dr. L. Doering, pers. comm.).

Recombinant human basic fibroblast growth factor (bFGF) stock solution, 10 ng/μl Resuspend lyophilized bFGF (Invitrogen) to a final concentration of 10 ng/μl in PBS (Invitrogen, cat. no. 14190-144) containing 0.1% (w/v) BSA. Store at −30◦ C.

Recombinant human epidermal growth factor (EGF) stock solution, 10 ng/μl Resuspend lyophilized EGF (Sigma) to a final concentration of 10 ng/μl in PBS (Invitrogen, cat. no. 14190-144) containing 0.1% (w/v) BSA. Store at −30◦ C.

Tumor sphere medium DMEM/F12 (Invitrogen) containing: 1× antibiotic-antimycotic (Wisent, cat. no. 450-115-EL; http://www.wisent.ca/) 1× hormone mix (N2 Supplement; Invitrogen, cat. no. 17502-048) 10 mM HEPES (Wisent, cat no. 330-050; http://www.wisent.ca/) 0.6% (w/v) glucose 60 μg/ml N-acetylcysteine (Sigma, cat no. A9165) 2% (w/v) NSF-1 (neural cell survival factor; Lonza, cat. no. CC-4323) Store for up to several weeks at 4◦ C Supplement the medium just before use with: 10 ng/ml LIF 20 ng/ml bFGF (see recipe for stock solution) 20 ng/ml EGF (see recipe for stock solution) COMMENTARY Background Information


When multipotent NSCs were isolated from the mammalian neuraxis more than a decade ago, culture conditions were developed that allowed embryonic EGF-responsive cells to proliferate as floating spheres (neurospheres), which could be easily manipulated for subsequent passage and differentiation (Reynolds and Weiss, 1992; Reynolds et al., 1992). Serum-free medium (SFM) allowed for the maintenance of an undifferentiated state, and the addition of saturating concentrations of bFGF and EGF (20 ng/ml) induced the proliferation of multipotent, self-renewing and ex-

pandable neural stem cells (Reynolds et al., 1992; Reynolds and Weiss, 1996). This neurosphere culture system and analysis procedure to identify NSCs has permitted in vitro characterization of these cells, but in a retrospective fashion, as the multipotential floating clusters of cells are inferred to have been derived from clonal expansion of a single NSC (Tropepe et al., 1999). Prospective study of this cell has been previously limited by lack of cell surface markers necessary for its isolation; recent reports show NSC enrichment using antibodies against the cell surface protein CD133 (Yin et al., 1997; Yu et al., 2002). Uchida and

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colleagues determined that this 120-kDa fivetransmembrane cell surface receptor of unknown function could effectively sort sphereforming cells from their non-sphere-forming counterparts in isolates of fetal human brain. Normal CD133+ human fetal brain cells not only efficiently form neurospheres in vitro, but also demonstrate the key stem cell properties of self-renewal and multilineage differentiation, and are capable of seamless lifelong engraftment and multilineage contribution to the mouse brain (Uchida et al., 2000). These findings represented the first evidence that the in vitro neurosphere-forming cell, when prospectively isolated, bore key stem cell properties both in vitro and in vivo. We applied the neurosphere culture assay to human brain tumors of different phenotypes, in order to select for stem cell growth and functionally characterize human brain tumor cell populations. Regardless of pathological subtype, within 24 to 48 hr of primary culture most brain tumors yielded a minority fraction of cells that demonstrated growth into clonally derived neurosphere-like clusters, termed tumor spheres. Tumor spheres are defined as clonally derived nonadherent colonies of cells derived from a single tumor stem cell. The remaining majority of tumor cells exhibited adherence, loss of proliferation, and subsequent differentiation, whereas tumor spheres remained nonadherent, continuing exclusively to self-renew and expand the tumor cell culture. From these cultures, the BTIC can then be exclusively isolated by fluorescence activated cell sorting for the neural precursor cell surface marker CD133 (Yin et al., 1997; Yu et al., 2002). Only the CD133+ brain tumor fraction contains cells that are capable of sphere formation and sustained self-renewal in vitro, and tumor initiation in NOD-SCID mouse brains (Singh et al., 2003, 2004). Therefore, our characterization of CD133+ BTICs lends support for the application of the cancer stem cell hypothesis to solid tumors, and our in vitro and in vivo BTIC models will provide the foundation for a brain cell hierarchy that may begin to organize the heterogeneity of brain tumors.

Critical Parameters and Troubleshooting There are many different methods of culturing brain tumor cells, and many variable applications of the neurosphere assay to human brain tumors. In culturing cells with stem cell properties in both normal and cancer tissues, there is no standardized protocol with respect

to growth factors, hormones, and their concentrations (Chaichana et al., 2006). In establishing human tumor sphere cultures, we have found that our tumor sphere medium and its components have been optimized for growth of healthy spheres with good cell viability and reliable stem cell frequency across different tumor subtypes. We recommend plating the cells at a high density (e.g., 2 × 105 cells per cm2 ), and we anticipate a large amount of cell death in the first few days of culture. This cell death can be attributed to both the elevated apoptotic activity of cancer cells and to the fact that the bulk tumor population will not survive in serum-free conditions, which does select for stem and progenitor cell growth. Another parameter within the tumor culture protocol that requires much testing and optimization is the enzyme digestion. Both the length of time for tumor digestion and the choice of enzyme/protease mix will influence the yield of tumor cells and their viability. Length of time for tumor specimen digestion must be judged based on each individual tumor, with larger and firmer specimens requiring longer digestion. Tumor pathological subtype may also influence this decision, as some benign brain tumors have a more extensive collagen- or fibrin-based framework, and some malignant tumors may harbor a greater degree of pre-existing tissue necrosis. Underdigestion will not yield the largest possible cell number, whereas overdigestion results in DNA lysis and increased amounts of stringy white fibers and cellular debris in the culture. In terms of choice of enzyme/protease mix for digestion, we have begun to favor the use of collagenase-based cocktails, due to the fact that CD133 is trypsin-sensitive (Fukuchi et al., 2004; Schwab et al., 2008), and this receptor may be cleaved during the digestion process. If the cells are not stringently treated with antitrypsin or are not allowed to re-equilibrate for a long enough time in tumor sphere medium post-digestion, the CD133 expression level on cell sorting may be underestimated. For sorting either with FACS or MBCS, it is critical to obtain a single-cell suspension, and neither normal neurospheres nor adherent tumor spheres are easily amenable to dissociation. Thus, we use a combination of very gentle mechanical trituration just prior to sorting, with filtering through cell-strainer caps into flow cytometry tubes, to prevent clumping during the sort. We also recommend treating the cells with chemical cell dissociation buffers, which may provide a more gentle alternative to mechanical dissociation.

Cancer Stem Cells


Supplement 11

Anticipated Results Depending on the size of the original tumor specimen, our tissue culture protocol should yield 1–100 × 106 viable tumor sphere cells from brain tumors of different subtypes, which then can be subjected to a panel of in vitro stem cell assays, including primary sphere formation assays, proliferation assays, limitingdilution assays, and differentiation assays. Our BTIC enrichment protocol should provide millions of CD133− brain tumor non-stem cells, and thousands to millions of CD133+ BTICs, depending on the clonogenic frequency and CD133 index of each tumor subtype. In general, the increased self-renewal capacity of the BTIC and the correlated CD133 index are highest from the most aggressive clinical samples of medulloblastoma and glioblastoma compared with low-grade gliomas. Both bead and FACS separation routinely yield average purities of 80% to 90% for the CD133+ cells and >99.5% purity for CD133− cells; thus, these methods should be considered as good enrichment methods, not methods for isolation to purity of BTICs.

Time Considerations Tumor dissociation and plating of cells into tumor sphere cultures typically takes 2 to 4 hr, and cell sorting can be completed within 4 hr.

Literature Cited Bonnet, D. and Dick, J.E. 1997. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat. Med. 3:730-737. Chaichana, K., Zamora-Berridi, G., CamaraQuintana, J., and Quiñones-Hinojosa, A. 2006. Neurosphere assays: Growth factors and hormone differences in tumor and nontumor studies. Stem Cells 24:2851-2857. Fukuchi, Y., Nakajima, H., Sugiyama, D., Hirose, I., Kitamura, T., and Tsuji, K. 2004. Potential human placenta-derived cells have mesenchymal stem/progenitor cell. Stem Cells 22:649-658. Reya, T., Morrison, S.J., Clarke, M.F., and Weissman, I.L. 2001. Stem cells, cancer, and cancer stem cells. Nature 414:105-111.

Reynolds, B.A. and Weiss, S. 1992. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707-1710. Reynolds, B.A., Tetzlaff, W., and Weiss, S.A. 1992. Multipotent EGF-responsive striatal embryonic progenitor cell produces neurons and astrocytes. J. Neurosci. 12:4565-4574. Reynolds, B.A. and Weiss, S. 1996. Clonal and population analyses demonstrate that an EGFresponsive mammalian embryonic CNS precursor is a stem cell. Dev. Biol. 175:1-13. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. 2009. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Schwab, K.E., Hutchinson, P., and Gargett, C.E. 2008. Identification of surface markers for prospective isolation of human endometrial stromal colony-forming cells. Hum. Reprod. 23:934-943. Singh, S.K., Clarke, I.D., Terasaki, M., Bonn, V.E., Hawkins, C., Squire, J., and Dirks, P.B. 2003. Identification of a cancer stem cell in human brain tumors. Cancer Res. 63:5821-5828. Singh, S.K., Hawkins, C., Clarke, I.D., Squire, J.A., Bayani, J., Hide, T., Henkelman, R.M., Cusimano, M.D., and Dirks, P.B. 2004. Identification of human brain tumor initiating cells. Nature 432:396-401. Tropepe, V., Sibilia, M., Ciruna, B.G., Rossant, J., Wagner, E.F., and van der Kooy, D. 1999. Distinct neural stem cells proliferate in response to EGF and FGF in the developing mouse telencephalon. Dev. Biol. 208:166-188. Uchida, N., Buck, D.W., He, D., Reitsma, M.J., Masek, M., Phan, T.V., Tsukamoto, A.S., Gage, F.H., and Weissman, I.L. 2000. Direct isolation of human central nervous system stem cells. Proc. Natl. Acad. Sci. U.S.A. 97:14720-14725. Yin, A.H., Miraglia, S., Zanjani, E.D., AlmeidaPorada, G., Ogawa, M., Leary, A.G., Olweus, J., Kearney, J., and Buck, D.W. 1997. AC133, a novel marker for human hematopoietic stem and progenitor cells. Blood 90:5002-5012. Yu, Y., Flint, A., Dvorin, E.L., and Bischoff, J. 2002. AC133-2, a novel isoform of human AC133 stem cell antigen. J. Biol. Chem. 277:2071120716.


3.3.10 Supplement 11


Human iPS Cell Derivation/Reprogramming

UNIT 4A.1

In-Hyun Park1 and George Q. Daley1, 2 1

Children’s Hospital Boston and Dana-Farber Cancer Institute, Harvard Medical School, Harvard Stem Cell Institute, Boston, Massachusetts 2 Brigham and Women’s Hospital, Howard Hughes Medical Institute, Boston, Massachusetts

ABSTRACT This unit describes a protocol for deriving induced pluripotent stem (iPS) cells from human fibroblast cells. Human fibroblast cells are infected with retroviral vectors expressing four transcription factors (Oct4, Sox2, Klf4, and Myc) and selected for 3 to 4 weeks under human embryonic stem (hES) cell culture conditions. iPS cell colonies are mechanically isolated using a dissection microscope and handled like hES cells thereafter. Human iPS cells share similarities with hES cells including the expression of pluripotency genes, and differentiation as embryoid bodies in vitro into three germ layers (EB) and in vivo C 2009 by John Wiley & as teratomas. Curr. Protoc. Stem Cell Biol. 8:4A.1.1-4A.1.8. Sons, Inc. Keywords: human induced pluripotent stem (iPS) cells r human embryonic stem (hES) cells r reprogramming r retroviral vectors

INTRODUCTION This unit describes a protocol for deriving induced pluripotent stem (iPS) cells from human fibroblasts. hES cells have the property of self-renewal and pluripotency that provides an unlimited resource for research and medical applications. Recently, terminally differentiated murine and human somatic cells were induced to become pluripotent stem (iPS) cells by use of a four-transcription-factor cocktail (Oct-4, Sox-2, Klf4, and Myc; Takahashi and Yamanaka, 2006; Takahashi et al., 2007; Yu et al., 2007; Park et al., 2008). In this unit, the production of retrovirus-expressing reprogramming factors, infection of human fibroblasts, and isolation of human iPS cells will be described. NOTE: The following tissue culture procedures are performed in a Class II biological hazard flow hood or a laminar-flow hood. NOTE: All procedures for producing the VSV-G pseudotyped retrovirus should be performed under BL2+ biosafety conditions (according to your Institute’s Safety Department). NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator.

PRODUCTION OF VSV-G PSEUDOTYPED RETROVIRUS This protocol is used for making retroviral vectors pseudotyped with the VSV-G (vesicular stomatitis virus G) envelope protein. The VSV-G pseudotyped retrovirus can be divided into aliquots and stored long-term at −80◦ C and then thawed and used to infect human fibroblasts.

BASIC PROTOCOL 1

Materials 293T cells (ATCC, cat. no. CRL11268) Fugene 6 (Roche Applied Science, cat. no. 1181509001) Current Protocols in Stem Cell Biology 4A.1.1-4A.1.8 Published online January 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc04a01s8 C 2009 John Wiley & Sons, Inc. Copyright

Manipulation of Potency

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DMEM DMEM/F12 (Invitrogen) pMIG-OCT4 (Addgene, clone 17225), pMIG-SOX2 (Addgene, clone 17226), pMIG-KLF4 (clone 17227), and pMIG-MYC (requested from Dr. Cleveland from the Scripps Research Institute) VSV-G (Addgene, clone 8454), and Gag-Pol (Addgene, clone 8455) 293T cell medium (see recipe) 10-cm dishes 0.45-μm filter 38.5-ml polyallomer centrifuge tube (Beckman, cat. no. 326823) Cryovials Additional reagents and equipment for determining the titer of the virus (Sastry et al., 2002; Tiscornia et al., 2006) Transfect 293T cells with plasmids using Fugene 6 1. One the day before transfection, split 293T cells (0.5 × 105 cells/cm2 ) into ten 10-cm dishes for each different virus at a confluency of 40%, aiming to have 70% to 80% confluency the next day. 2. For each 10-cm dish, add 20 μl of Fugene to 300 μl DMEM and incubate the mixture 15 min at room temperature. Add 2.5 μg of pMIG vector, 2.25 μg of Gag-Pol, and 0.25 μg of VSV-G vector and incubate for 15 min. For ten 10-cm dishes, multiply the amounts by 10 to have 3 ml of DMEM with 200 μl Fugene, 25 μg pMIG vector, 22.5 μg Gap-Pol and 2.5 μg VSV-G. Use polystyrene tubes for maximum transfection.

3. While the Fugene/plasmid mixture of step 2 is incubated, aspirate old medium from 293T cells and add 9 ml of new 293T medium. 4. Add 7 ml of 293T cell medium into 3 ml of Fugene/plasmid mixture (for ten 10-cm dishes from step 2) to make a total of 10 ml, mix well by pipetting and add 1 ml of each into each 10-cm dish of 293T cells in 9 ml medium. 5. Place the transfected 293T cells into BL2+ incubator. After 293T cells are transfected with retroviral vectors, they need to be treated as BL2+ hazardous biomaterial.

Concentrate VSV-G pseudotyped retrovirus 6. Three days after transfection (do not change medium during 3-day incubation period), collect and filter the viral supernatant using 0.45-μm filter into a 38.5-ml polyallomer centrifuge tube. 7. Weigh the supernatant tube to make a balance tube for ultracentrifugation. 8. Centrifuge the supernatant 90 min at 70,000 × g, 4◦ C. 9. Remove the supernatant. A white pellet should be visible when a polyallomer centrifuge tube is used.

10. Add 1 ml DMEM/F12 and flick the tubes before storing overnight at 4◦ C to dissolve the pellet. 11. Next day, mix virus by pipetting up and down slowly, aliquot ∼100 to 200 μl of virus into cryovials and store at −80◦ C for long-term storage. Human iPS Cell Derivation/ Reprogramming

12. Determine the titer of the virus following published protocols (Sastry et al., 2002; Tiscornia et al., 2006).

4A.1.2 Supplement 8


INFECTION OF FIBROBLASTS AND ISOLATION OF iPS CELLS This protocol is used to infect fibroblasts with the virus, and to isolate iPS cell colonies from them. During and after isolation, human iPS cells show similar colony morphology and require the same culture conditions as human ES cells. Therefore, it is highly recommended that scientists wishing to isolate and maintain human iPS cells first become skilled in handling hES cells, including the mechanical picking and passaging of colonies (UNIT 1C.1), or undergo training directly on iPS cell culture from experienced investigators.

BASIC PROTOCOL 2

Materials Human fibroblasts, acquired from skin biopsy (refer to UNIT 1C.7; split fibroblasts when they reach 70% confluency) Human fibroblast medium (see recipe) Protamine sulfate (see recipe) Retroviral supernatants bearing the appropriate plasmids (Basic Protocol 1) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Mediatech, cat. no. 21-040-CV) MEF (mouse embryonic fibroblasts), CF-1 strain, irradiated (Global Stem, cat. no. GSC-6001G) MEF medium (see recipe) 0.1% (w/v) gelatin (see recipe) 0.05% trypsin/EDTA hESC medium (see recipe) Gelatin-coated 12-well plate preplated with MEFs at a density of 1 × 104 cells/cm2 Collagenase IV (see recipe) Gelatin-coated 6-well plate preplated with MEFs Freezing medium (see recipe) Liquid nitrogen 6-well plate 10-cm dish Dissection microscope 20- and 1000-μl pipets 21-G needle or cell lifter (Corning, cat. no. CT-3008) 15-ml conical tube Cryovials −80◦ C freezer Infect human fibroblasts with retrovirus 1. At a time point 8 to 12 hr prior to infection, plate 1 × 105 human fibroblasts obtained from a skin biopsy in one well of a 6-well plate. 2. Aspirate medium to remove dead cells and add 2 ml of fresh human fibroblast medium. Add protamine sulfate at a final concentration of 5 μg/ml. 3. Into the fibroblast culture, add aliquots of the four retroviral supernatants (Basic Protocol 1) at a multiplicity of infection (MOI) of 5 for each virus. This step and those following must be performed under BL2+ safety conditions.

4. One day after infection, remove the viral supernatant, wash three times, each time with 3 ml PBS, and add 3 ml human fibroblast medium. 5. After 2 more days, replenish dish with 3 ml human fibroblast medium. 6. Four days after infection, plate 1 × 104 /cm2 MEFs in MEF medium on a 10-cm dish coated with 0.1% gelatin. Incubate until the next day.



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Figure 4A.1.1 Identification of iPS colonies among heterogeneous types of colonies arising from infected human fibroblasts. Examples of colonies that should yield stable iPS colonies (A, B) and examples of colonies that do not (C, D). Silencing of retroviral gene expression in iPS colony (E, F). (A) iPS colony with a morphology comparable to hES cells. (B) iPS colony that is surrounded by outgrowth of infected fibroblasts. (C) Non-iPS colony from transformed cells. (D) Non-iPS colony from transformed cells. (E) iPS cell colony in the middle of field (arrow). (F) A reprogrammed iPS colony in E shows no fluorescence due to silencing of the infected retrovirus (arrow). Scale bar = 100 μm.

7. Five days after infection, incubate the infected fibroblasts with 1 ml 0.05% trypsin/EDTA for 3 min at 37◦ C. Stop the trypsinization with 11 ml human fibroblast medium and place all cells into one 10-cm dish preplated with MEFs (prepared in step 6). 8. Seven days after infection remove human fibroblast medium and add 10 ml hESC medium (containing KOSR and 10 ng/ml of bFGF). 9. Replenish the cells with 10 ml hESC medium every day and observe the cells for any sign of colony formation. Typically, sometime around three weeks after infection, various types of colonies will appear, as exemplified in Figure 4A.1.1.

10. Under a dissection microscope and using a 20-μl pipet, pick a colony that shows a morphology similar to hES cells and put the colony into one well of a 12-well plate that has been preplated with MEFs at a density of 1 × 104 cells/cm2 on 0.1% gelatin-coated wells. Fully reprogrammed iPS cells infected with GFP-containing viruses will lack GFP expression, as visualized under a fluorescence microscope (Fig. 4A.1.1F). Absence of GFP expression, a consequence of transcriptional silencing of retroviruses, is specific to embryonic or pluripotent cell types, and can be used as a surrogate marker to identify human iPS cells.

Human iPS Cell Derivation/ Reprogramming

Maintain and store human iPS cells Isolated clones of human iPS cells can be treated and maintained in a similar way as human ES cells. Please refer to the detailed protocol UNIT 1C.1.

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11. When iPS cell colonies grow and start to touch other colonies, wash plate with 1 ml of DMEM/F12, add 0.5 ml collagenase IV, and incubate 10 min at 37◦ C. 12. Using appropriate tools (e.g., 21-G needle, or cell lifter), cut colonies into small pieces, detach pieces from the plate, collect with a 1000-μl pipet (similar to mechanical splitting of established human ES cells; UNIT 1C.1) into a 15-ml conical tube and centrifuge for 4 min at 200 × g, room temperature. Remove the supernatant and add 5 ml of fresh DMEM/F12 followed by centrifuging 4 min at 200 × g, room temperature. 13. Remove the supernatant. 14. For passaging of iPS cells, resuspend pieces of colonies from step 13 in 2 ml fresh hESC medium and transfer into one well of a gelatin-coated 6-well dish precoated with MEFs. Repeat steps 10 to 14 to expand iPS cells. The split ratio used for the cells depends on cell density (ratio is usually 1:3 to 1:6) When you have more than three wells of iPS cells in a 6-well plate, freeze down cells before starting further analysis of iPS cells.

Freeze cells 15. For freezing iPS cells, resuspend colonies from step 13 in 0.5 ml fresh hESC medium, add the same amount of 2× freezing medium dropwise and mix by gently pipetting up and down. Aliquot cells into cryovials and store overnight at –80◦ C. Transfer to liquid nitrogen next day. REAGENTS AND SOLUTIONS For culture recipes and steps, use sterile tissue culture–grade water. For other purposes, use deionized, distilled water or equivalent in recipes and protocol steps. For suppliers, see SUPPLIERS APPENDIX.

293T cell medium DMEM containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C Collagenase IV, 10× Dissolve collagenase IV (Invitrogen) at 10 mg/ml in DMEM/F12 (Invitrogen). Filter using 0.22-μm filter. Divide into 0.5- to 1.5-ml aliquots and store up to 1 year at −20◦ C. Before splitting hES or iPS cells, dilute 10× stock solution in DMEM/F12 to make a 1× working stock.

Freezing medium, 2× Make a solution containing 20% dimethyl sulfoxide (DMSO), 60% FBS, and 20% human ES cell (hESC) medium (see recipe). Store up to 1 month at 4◦ C. Gelatin, 0.1% (w/v) Dissolve 0.5 g of gelatin (from porcine skin) in 500 ml distilled water and autoclave. Store indefinitely at room temperature. To gelatinize plates: Prior to addition of MEFs, coat all dishes or wells with enough 0.1% (w/v) gelatin solution to cover the surface. Remove gelatin after 5 min.



Supplement 8

Human embryonic cell (hESC) medium DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (KOSR; Invitrogen) 10 mM non-essential amino acids 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) 50 mM 2-mercaptoethanol 10 ng/ml bFGF (see recipe) Store up to 1 week at 4◦ C Human fibroblast medium MEM-alpha containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C MEF medium DMEM containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine 1× penicillin/streptomycin (add from 200× stock, e.g., Invitrogen) Store up to 4 weeks at 4◦ C Protamine sulfate, 1000× Dissolve protamine sulfate (Sigma-Aldrich) at 5 mg/ml in phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; Mediatech, cat. no. 21-040-CV). Filter using a 0.22-μm filter and store up to 1 year at 4◦ C for future use.

Recombinant human basic fibroblast growth factor (bFGF) Resuspend lyophilized bFGF (PeproTech) to a final concentration of 10 μg/ml in CMF-PBS containing 0.1% (w/v) bovine serum albumin (BSA) and 1 mM DTT. Store at −80◦ C according to manufacturer’s instructions.


Human iPS Cell Derivation/ Reprogramming

Human embryonic stem cells provide a valuable resource for research and regenerative medicine. However, human ES cells isolated to date are not matched to individual patients, and thus allow for generic studies but are limited in their relevance to specific diseases or treatments. Attempts to generate patientspecific stem cells include somatic cell nuclear transfer (SCNT), somatic cell fusion with pluripotent cells, direct cultural adaptation of germ cells, and direct reprogramming of somatic cells with defined factors (Jaenisch and Young, 2008). Each of these approaches has specific advantages and limitations. Since the report by Yamanaka’s group that mouse embryonic and adult tail-tip fibroblasts can be reprogrammed to become ES cell–like pluripo-

tent cells by expression of four transcription factors (Oct4, Sox2, Klf4, c-Myc), a similar strategy has been used to isolate human iPS cells. Our laboratory and Yamanaka’s group successfully isolated human iPS cells from embryonic and adult human fibroblasts using the same four transcription factors (Takahashi et al., 2007; Park et al., 2008). A different combination of factors (Nanog and Lin28 in place of Klf4 and Myc) also produced human iPS cells (Yu et al., 2007). Yamanaka’s reprogramming strategy also works even without Myc, although at lower efficiency, to produce iPS cells from both murine and human fibroblasts (Nakagawa et al., 2008). The isolation of murine iPS cells has been facilitated by using fibroblasts that carry endogenous selectable markers. Fibroblasts from

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mice that express neomycin- or puromycinresistance genes under the control of the promoters for Fbx15, Oct4, or Nanog loci have been infected with four reprogramming factors, selected with neomycin or puromycin, and successfully reprogrammed to become iPS cells (Takahashi and Yamanaka, 2006; Maherali et al., 2007; Okita et al., 2007; Wernig et al., 2007). For most human fibroblasts, such endogenous selection systems do not exist, and thus selection has been based mostly upon colony morphology alone. By selecting colonies that show similar morphology to human ES cells, human iPS cells can be readily selected from a morphologically diverse background of colonies that arise when human fibroblasts are infected with several retroviruses. Although reprogramming human fibroblasts into pluripotent cells will provide an alternative to human ES cells for certain research and clinical applications, current methods of generating iPS cells employ retroviral vectors that integrate into the fibroblast genome, and thus the resulting cells are potentially tumorigenic. Developing virus-free methods is desirable. Excision of ectopic Myc by Cremediated recombination has been shown to reduce the tumor formation potential of iPS cells (Hanna et al., 2007; Shi et al., 2008). The G9a inhibitor BIX01294 can replace Oct4, and only two factors are sufficient to make iPS cells from murine neuronal stem cells (Shi et al., 2008), suggesting that the right combination of excisable viruses and chemicals may provide a method to make iPS cells that lack persistent viral integration.

Critical Parameters and Troubleshooting Before starting to reprogram human fibroblasts, there are three important factors that determine success: the quality of the virus, MEFs, and target fibroblasts. When a new VSV-G pseudotyped virus is made and stored at –80◦ C, be sure to determine the titer of the virus, because it is essential to have a highquality, high-titer virus. If the titer is 100 ng/μl RNA for RT-PCR. You can stop the experiment at this step. Purified RNA samples should be stored at −80◦ C.

Perform reverse transcription 16. Prepare 20 μl of reaction mixture by mixing the reagents listed below: 4 μl 5× reverse transcription buffer (from ReverTra Ace kit) 2 μl 10 mM dNTPs (from ReverTra Ace kit) 1 μl ribonuclease inhibitor (from ReverTra Ace kit) 1 μl ReverTra Ace (reverse transcriptase; from ReverTra Ace kit) 1 μl 10 μM oligo dT20 primer (from ReverTra Ace kit) 1 μg DNase-treated total RNA (step 14) Nuclease-free water up to 20 μl. You should prepare reactions containing no reverse transcriptase as negative controls for each sample.

17. Incubate the mixture in thermal cycler at the condition as follows:

60 min at 42◦ C 5 min at 95◦ C Indefinitely at 4◦ C. You can stop the experiment at this step. cDNA samples should be stored at −20◦ C or lower.

Amplify the products by PCR 18. Prepare 25 μl of PCR mixture by mixing the reagents listed below in a 0.2-ml PCR reaction tube: 2.5 μl 10× ExTaq buffer (from ExTaq kit) 2 μl 2.5 mM dNTPs (from ExTaq kit) 0.5 μl 10 μM forward primer (Figure 4A.2.3) 0.5 μl 10 μM reverse primer (Figure 4A.2.3) 0.5 μl 5 U/μl ExTaq DNA polymerase (from ExTaq kit) 1 μl cDNA sample (step 17) 1.25 μl of DMSO (optional, depends on primer sets) Nuclease-free water up to 25 μl.



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19. Carry out PCR according to the conditions listed in Figure 4A.2.3. PCR conditions, particularly the number of cycles, may differ among different thermal cyclers. It is necessary to experiment to find the optimal conditions.

20. After finishing PCR, analyze the by electrophoresis on a 2% agarose gel in 1× TAE buffer using a standard protocol (e.g., Voytas, 2000). SUPPORT PROTOCOL 4

IMMUNOCYTOCHEMISTRY FOR PLURIPOTENT CELL MARKERS The expression of pluripotent stem cells marker can be confirmed not only by RT-PCR (Support Protocol 3) but also by immunocytochemistry. Some surface antigens specifically expressed in pluripotent cells such as SSEAs and TRAs were identified by analyses of human embryonic carcinoma (EC) and ES cells. See Figure 4A.2.2 for examples of immunohistochemistry results.

Materials Human iPS cells (Basic Protocol 1) 6-well plates seeded with mitomycin C–treated feeder cells (Support Protocol 1) hES medium (see recipe) Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CMF-DPBS containing 10% (v/v) formalin CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum (or donkey serum), and 0.2% (v/v) Triton X-100 (omit Triton if staining surface antigens) Primary antibodies against desired ES markers (perform all dilutions in CMF-DPBS containing 1% v/v bovine serum albumin): Anti-Nanog goat polyclonal (R&D Systems, cat. no. AF1997; use at 1:20 dilution) Anti-SSEA-1 mouse IgM (Developmental Studies Hybridoma Bank, cat. no. MC480; use at 1:5 dilution) Anti-SSEA-3 rat IgM (Developmental Studies Hybridoma Bank cat. no. MC631; use at 1:5 dilution) Anti-TRA-1-60 mouse IgM (Chemicon, cat. no. MAB4630; use at 1:50 dilution) Anti-TRA-1-81 mouse IgM (Chemicon, cat no. MAB4381; use at 1:50 dilution) Secondary antibody against IgG or IgM of species in which primary antibody was raised, labeled with Alexa Fluor 488 or Alexa Fluor 546; use at 1:500 dilution in CMF-DPBS containing 1% (w/v) bovine serum albumin 10 mg/ml Hoechst 33342 (H3570, Invitrogen) 1. To prepare cells for immunostaining, seed about 100 to 200 clumps of human iPS cells in hES cell medium in each well of a 6-well plate containing mitomycin-treated SNL feeder cells and incubate for 5 to 7 days prior to fixation.

Fix cells and block nonspecific binding 2. Prior to fixation, aspirate the medium, and wash with 2 ml of CMF-DPBS. 3. Remove CMF-DPBS, add 2 ml of CMF-DPBS containing 10% formalin, and fix the cells by incubating for 10 min at room temperature. 4. After fixation, wash the cells once with 2 ml of CMF-DPBS.

Generation and Characterization of Human iPS Cells

5. Aspirate CMF-DPBS and add 2 ml of CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum, and 0.2% (v/v) Triton X-100. Incubate 45 min at room temperature.



Omit Triton X-100 when staining for surface antigens. Triton X-100 is not necessary for immunostaining of surface antigens. Treatment with Triton X-100 is required only for anti-Nanog antibody. For anti-Nanog antibody, substitute normal donkey serum for normal goat serum because anti-Nanog antibody was raised in goat.

Treat cells with primary and secondary antibodies 6. After blocking procedure, incubate the cells 1 ml of primary antibody at the appropriate dilution in CMF-DPBS containing 1% bovine serum albumin, overnight at 4◦ C. Other antibodies should work. Determine the optimal dilution.

7. Wash the cells three times each for 5 min with CMF-DPBS. 8. Add 1 ml of secondary antibody conjugated with Alexa Fluor 488 or 546 to the sample at the appropriate dilution in CMF-DPBS containing 1% bovine serum albumin supplemented with 1 μg/ml of Hoechst 33342 (added from 10 mg/ml Hoechst stock solution), and incubate for 45 min at room temperature in the dark. 9. Wash out secondary antibody with 2 ml CMF-DPBS three times, each time for 5 min. 10. Observe the cells with a fluorescent microscope equipped with the appropriate filters.

ASSESSING PLURIPOTENCY BY IN VITRO DIFFERENTIATION OF iPS CELLS BY EMBRYOID BODY FORMATION

SUPPORT PROTOCOL 5

Embryoid body formation is one of the easiest procedures for in vitro differentiation of ES cells. This also can be applied for differentiation of iPS cells. Our protocol consists of a primary floating culture for 8 days. After 8 days of floating culture, transfer the cells to gelatin-coated plates to induce further differentiation. After embryoid body formation, differentiation should be confirmed by immunocytochemistry for differentiated markers. Other procedures such as RT-PCR (Support Protocol 3) are also suitable for determination of pluripotency and/or differentiation.

Materials 10 mg/ml HEMA-MMA (see recipe) Growing human iPS cells (Basic Protocols 1 and 2) at 80% to 90% confluency in 60-mm dish Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium containing no bFGF (see recipe) CMF-PBS containing 10% (v/v) formalin (Sigma) CMF-PBS containing 1% (w/v) bovine serum albumin, 5% (v/v) normal goat serum (or donkey serum), and 0.2% (v/v) Triton X-100 Primary antibodies against desired ES markers for immunohistochemistry (perform all dilutions in CMF-PBS containing 1% v/v bovine serum albumin): Anti-α-fetoprotein mouse IgG (R&D Systems, cat. no. MAB1368; use at 1:100 dilution) Anti-α-smooth muscle actin mouse IgG (Dako, cat. no. N1584; use at 1:500 dilution) Anti-βIII-tubulin mouse IgG (Chemicon, cat. no. CB412; use at 1:100 dilution) Secondary antibody: anti-mouse IgG labeled with Alexa Fluor (use at 1:500 dilution in CMF-DPBS containing 1% w/v bovine serum albumin) 10 mg/ml Hoechst 33342 solution (Invitrogen)



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100-mm tissue culture dish Sterile disposable cell scraper 15-ml conical centrifuge tubes Gelatin-coated 6-well culture plate (see recipe) Additional reagents and equipment for immunohistochemistry (Support Protocol 4) Establish suspension culture 1. Place 5 ml of 10 mg/ml of HEMA-MMA in a 100-mm dish, and incubate at room temperature in a hood with the dish covered with foil until the solution dries up (3 to 5 days). 2. Wash the iPS cells in 60-mm dish once with 4 ml CMF-DPBS. 3. And add 0.5 ml of CTK solution and return the dish to the 37◦ C incubator. 4. After 5 min incubation, wash twice with 4 ml of CMF-DPBS to remove the CTK solution and detached feeder cells. 5. Add 4 ml hES medium without bFGF to the dish. 6. Detach iPS colonies from the dish by using cell scraper. Collect the cell clumps to a 15-ml conical tube. Do not break up the colonies; larger colonies can form embryoid bodies effectively.

7. Add another 5 ml of hES medium without bFGF and transfer the cell suspension to the HEMA-coated 100-mm dish from step 1. 8. Incubate 2 days in humidified 37◦ C, 5% CO2 incubator. 9. To change the medium, collect the cell suspension into a 15-ml conical tube and let sit it for 5 min at room temperature. 10. Remove the supernatant (∼8 ml) carefully, then add 8 ml of fresh hES medium without bFGF and return the suspension to a HEMA-coated dish prepared as in step 1. Change the medium every other day.

Set up attached culture 11. Collect the iPS cell suspension into a 15-ml conical tube, and let sit for 5 min at room temperature. Remove the supernatant and resuspend the cells in 12 ml of hES medium without bFGF. 12. Transfer 2 ml of cell suspension into wells of a gelatin-coated 6-well culture plate, and incubate at 37◦ C, 5% CO2 . 13. Change the medium every other day. 14. After 8-day attached culture, perform immunocytochemistry for differentiated cell markers (see Support Protocol 4). We routinely observe the expression of α-fetoprotein for endoderm, α–smooth muscle actin for mesoderm, and βIII-tubulin for ectoderm. Other antibodies and markers may also be used for this purpose. SUPPORT PROTOCOL 6 Generation and Characterization of Human iPS Cells

ASSESSING PLURIPOTENCY BY IN VIVO DIFFERENTIATION BY TERATOMA FORMATION Teratoma formation is another well known, important test of pluripotency. In general, mouse ES and iPS cells can produce teratomas easily. However, it is hard to form tumors derived from either human ES or iPS cells by subcutaneously injection into immunodeficient mice, including NOD-SCID mice. Therefore, in this protocol we inject



stem cells into testes of SCID mice. This change improves the efficiency of tumor formation to more than 80%.

Materials 10 mM Y-27632 (Wako, cat. no. 253-00513) Growing iPS cells (Basic Protocols 1 and 2) at 80% to 90% confluency in 60-mm dish Dulbecco’s phosphate-buffered saline without calcium and magnesium (CMF-DPBS: Nacalai Tesque, cat. no. 14249-95) CTK solution (see recipe) hES medium (see recipe) DMEM/F12 medium (e.g., Invitrogen) supplemented with 10 μM Y-27632 1.2% tribromoethanol (Avertin): dissolve 2.5 g tribromoethanol in 5 ml butanol, then add 200 ml distilled water; store at 4◦ C in the dark SCID mice, (7- to 8-weeks, male) 70% ethanol CMF-PBS containing 10% formalin Sterile disposable cell scrapers 15-ml conical centrifuge tubes Centrifuge Hamilton syringe 25-G to 26-G needle (Terumo) Suture needle with thread Additional reagents and equipment for intraperitoneal injection (Donovan and Brown, 2006a) and euthanasia of the mouse (Donovan and Brown, 2006b), paraffin embedding and sectioning of tissue, and hematoxylin/eosin staining of tissue sections (UNIT 2A.5) NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to government regulations for the care and use of laboratory animals.

Prepare cell suspension 1. Add 10 μM Y-27632 (from 10 mM stock) to the medium of a confluent culture of iPS cells, and incubate at 37◦ C for at least 1 hr. Y-27632 is a specific inhibitor for p160-Rho-associated coiled-coil kinase (ROCK).

2. Wash the cells with 4 ml of CMF-DPBS, and add 0.5 ml of CTK solution. Incubate ∼5 min at room temperature. After treatment with Y-27632, the cells may become less detachable. In such cases, you can treat the cells with CTK solution for longer period of time (∼10 min).

3. Wash out CTK solution and detached feeder cells with 4 ml of CMF-DPBS, twice, and add 4 ml of hES medium. 4. Detach iPS cells from the dish with a cell scraper, and break the colonies into small clumps by pipetting up and down several times. 5. Collect the cell suspension to a 15-ml conical tube, and centrifuge 5 min at 200 × g, room temperature. 6. Aspirate the supernatant, and resuspend cells in 300 to 500 μl of DMEM/F12 supplemented with 10 μM of Y-27632. Manipulation of Potency


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Inject cells 7. Inject 0.8 ml of 1.2% tribromoethanol solution intraperitoneally (Donovan and Brown, 2006a) into SCID mouse (0.12 ml for 10 g weight). 8. Wash the lower abdominal/groin area with 70% ethanol. 9. Dissect out the testes and remove from the body. Dissect the lower abdominal/groin area and withdraw the inguinal canal and then the testes. Leave spermatic cord intact. 10. Inject 30 μl of iPS cell suspension into a testes, under the capsule, using a Hamilton syringe and a 25-G to 26-G needle, as gently as possibly. 11. Return the testes to the original interperitoneal location, and close the incision with stitches. Return mouse to colony within 2 hr. 12. About 3 months later, observe the mice for teratoma formation (Fig. 4A.2.2K). Mice may appear to be pregnant, indicating the presence of a teratoma.

Dissect the tumors 13. Euthanize mice (Donovan and Brown, 2006b) bearing teratomas and dissect out the tumors. 14. Fix the tumors in ∼50 ml of CMF-DPBS containing 10% formalin and incubate overnight at room temperature with agitation. 15. After fixation, embed the tumor in paraffin. 16. Slice the tumor into 4- to 5-μm sections and mount on slides. 17. Stain the sections with hematoxylin and eosin using a standard protocol (e.g., UNIT 2A.5). 18. Examine the entire set of sections for a tumor, scoring for the presence of derivatives of all three germ layers such as cartilage, pigmented epithelium, and gut-like epithelium (see Fig. 4A.2.2M). If the tumor contains derivatives of all three germ layers, the iPS cell line is pluripotent.


10% FBS medium DMEM (e.g., Invitrogen) containing: 10% fetal bovine serum (FBS) 50 U/ml penicillin 50 μg/ml streptomycin To prepare 500 ml of 10% FBS medium, mix 50 ml FBS and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week.


For Plat-E cells (see Support Protocol 2), add 1 μl of 10 mg/ml puromycin stock (see recipe) and 10 μl of 10 mg/ml blasticidin S stock (see recipe) to 10 ml of 10% FBS medium.



293FT medium DMEM (e.g., Invitrogen) containing: 10% fetal bovine serum (FBS) 2 mM L-glutamine 1 × 10−4 M nonessential amino acids 1 mM sodium pyruvate 50 U penicillin 50 μg/ml streptomycin 0.5 mg/ml G418 To prepare 500 ml of the medium, mix 50 ml of FBS, 5 ml of 200 mM (100×) L-glutamine, 5 ml 100× nonessential amino acids, 5 ml of 100 mM sodium pyruvate, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week. Just before use, add 100 μl of 50 mg/ml G418 to 10 ml 293FT medium.

Blasticidin S stock solution Dissolve blasticidin S hydrochloride (Funakoshi Chemical Company; http://www. funakoshi.co.jp) in distilled water at 10 mg/ml and sterilize through a 0.22-μm filter. Aliquot and store at −20◦ C.

CTK solution 5 ml 2.5% (w/v) trypsin 5 ml 1 mg/ml collagenase IV 0.5 ml 0.1 M CaCl2 10 ml Knockout Serum Replacement (KSR; Invitrogen) 30 ml distilled water Store up to 1 month at −20◦ C Do not repeat freeze/thaw cycles DAP213 solution To 5.37 ml hES medium (see recipe) add: 1.43 ml DMSO 1 ml 10 M acetamide 2.2 ml of propylene glycol Store up to 1 month at −80◦ C Gelatin coating of culture vessels Dissolve 1 g of gelatin powder (Sigma, cat. no. G-1890) in 100 ml of distilled water, autoclave, and store at 4◦ C as the 10× gelatin stock solution. To prepare 0.1% (1×) gelatin solution, thaw the 10× gelatin stock in a microwave and/or autoclave, then add 50 ml of the 10× solution to 450 ml of distilled water. Filter the solution with a 0.22-μm filter unit and store at 4◦ C. To coat culture dishes, add appropriate volume of 0.1% (1×) gelatin solution to cover the entire area of the dish bottom. For example, 1, 3, or 5 ml of gelatin solution is used for a 35-, 60-, or 100-mm dish, respectively. Incubate the dishes for at least 30 min at 37◦ C in a sterile environment. Before using, aspirate off the excess gelatin solution. Gelatin stock is prepared as 10× concentration (1% w/v) stocks.

hES medium DMEM/F12 medium containing: 20% Knockout Serum Replacement (KSR) 2 mM L-glutamine Current Protocols in Stem Cell Biology

Manipulation of Potency continued


1 × 10−4 M nonessential amino acids 1 × 10−4 M 2-mercaptoethanol 50 U penicillin 50 μg/ml streptomycin To prepare 500 ml of the medium, mix 100 ml KSR, 5 ml of 200 mM (100×) L-glutamine, 5 ml 100× nonessential amino acids, 1 ml 2-mercaptoethanol, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM/F12. Add 200 μl of 10 μg/ml bFGF into 500 ml of the medium just before use. For differentiation experiments (e.g., Support Protocol 5), do not add bFGF. Store at 4◦ C up to 1 week. All abovementioned components are available from Invitrogen. Primate ES cell medium from ReproCELL (http://www.reprocell.net/) may be used as an alternative.

Mitomycin C, 0.4 mg/ml Dissolve 2 mg of mitomycin C (Kyowa Hakko Kirin; http://www.kyowa-kirin.co.jp/ english/) in 5 ml of CMF-DPBS (Nacalai Tesque, cat. no. 14249-95). Store up to 1 month at −20◦ C in the dark. CAUTION: Because of its toxicity, the solution must be treated exclusively in a safety cabinet with gloves and lab coat and disposed of in accordance with the rules each institution stipulates.

Poly(hydroxyethyl methacrylate-co-methyl methacrylate; HEMA-MMA), 10 mg/ml Add 0.3 g of HEMA-MMA (Sigma, cat. no. P-3932) to a tube containing 30 ml ethanol. Incubate at 37◦ C overnight with agitation. Prepare fresh for each experiment.

Puromycin Dissolve puromycin (Sigma, cat. no. P-8833) in distilled water at 10 mg/ml and sterilize through a 0.22-μm filter. Divide into aliquots and store up to 1 month at −20◦ C.

SNL medium DMEM (e.g., Invitrogen) containing: 7% fetal bovine serum (FBS) 2 mM L-glutamine 50 U penicillin 50 μg/ml streptomycin To prepare 500 ml of the medium, mix 35 ml FBS, 5 ml 200 mM (100×) Lglutamine, and 2.5 ml of 100× penicillin/streptomycin (containing 10,000 U penicillin and 10,000 mg/ml streptomycin). Make up to 500 ml with DMEM. Store at 4◦ C up to 1 week. This medium is used for fibroblasts and PLAT-E cells.



Although it is commonly known that nuclei of differentiated cells can be reprogrammed back to embryonic states by means of nuclear transfer into oocytes or fusion with ES cells, the mechanism of inducing nuclear reprogramming has yet to be revealed. The fact that

somatic cells can be reprogrammed by fusion with ES cells implies that ES cells contain factors that can induce reprogramming. We hypothesized that factors which play important roles in ES cells also play pivotal roles in induction of nuclear reprogramming. Pluripotency and tumor-like proliferation are



the most exquisite properties of ES cells. Three transcription factors—Oct3/4, Sox2, and Nanog—have been found to be essential in the maintenance of pluripotency in both early embryos and ES cells. While a handful of laboratories have demonstrated that several tumor-related gene products, such as ERas, c-myc, and Stat3, contribute to long-term maintenance of ES cells in culture, we have identified several genes that are specifically expressed in ES cells by analyzing expressed sequence tag (EST) databases. After selecting the most promising 24 gene products as candidates for potential factors that could induce reprogramming, we narrowed these to four transcription factors (Oct3/4, Sox2, Klf4, and c-myc) that have been shown to convert fibroblasts back to pluripotent state. The identification of these factors was an important breakthrough that has revealed a mechanism of nuclear reprogramming and let us create pluripotent cells directly from skin biopsy specimens. One year later, other groups succeeded in generating iPS cells from human somatic cells. Recently, two research groups have reported that various disease-specific iPS cells from a patient’s own somatic cells have been successfully reprogrammed (Dimos et al., 2008; Park et al., 2008b). Now, iPS cell technology can be used in conjunction with or in place of ES cell technology to shed light on understanding pathogens, in drug discovery, and most of all, to develop regenerative medicine applications. Encouraging broad use of iPS cell technology will facilitate the development of practical applications. These protocols should provide guidance to scientists who share our objectives.

Troubleshooting In some cases, lentivirus is toxic to fibroblasts. Depending on the different cell lines, lentiviral transduction may lead to loss or growth arrest of fibroblasts due to their sensitivity to the virus. As some fibroblasts are more vulnerable to lentivirus than common cells, they should be treated with a double dilution of the virus-containing supernatant in fresh medium or by shortening the exposure time from overnight to 5 hr. For our purposes, the expression of mouse Slc7a1 gene is sufficient for generation of iPS cells despite lower infection efficiency. When no ES-like (iPS) colonies appear in fibroblast cultures after introduction of the four factors, the following causes should be considered. First, the titer of retrovirus may be too

low. Transduction efficiencies of retroviruses for reprogramming factors are critical for iPS cell colony formation as described above. The retrovirus must be prepared fresh for every experiment. Do not use frozen stock retroviruses because freezing causes reduction of the titer. Growth properties of the fibroblasts are also important for iPS cell generation. Efficiency of retroviral transduction is markedly reduced when senescent fibroblasts are used for transduction. We strongly recommend banking stocks of fibroblasts at early passages and using fresh fibroblasts of early passage for iPS cell production. The number of cells that are plated onto SNL feeder cells after retroviral transduction is important. Overgrowth of fibroblasts might make cells peel off from the edge of the dish like a sheet, inhibiting formation of iPS cell colonies. Although this may be overcome by reducing the cell number, too small a number of cells could lead to no colony appearance. The optimal conditions differ for each individual cell type. We recommend that you seed at least in two or three different dishes with different densities when first plating the transduced cells. In addition, the qualities of feeder cells are crucial not only for generation of iPS cells, but also for maintenance of them. If feeder cells are too old, cells may peel off the substrate during the reprogramming or maintenance. SNL feeder cells more than 3 days after mitomycin C treatment cannot survive the stimulation by bFGF in hES medium (as bFGF may have a toxic effect on older feeder cells). It is recommended that SNL feeder cells be used within 3 days after inactivation. Some problems may arise after iPS cells are generated. For example, iPS cells can change characteristics and potential, depending on the line, with long-term culture. Human iPS cells, like human ES cells, may become adapted in a long-term culture. We recommend that large amounts of iPS cell stocks be frozen at early passages to support long-term experimentation. iPS cells are relatively unstable during early passage period so that spontaneous differentiation in daily culture may also happen. When the number of differentiated colonies increases, select undifferentiated colonies and transfer them by aspiration to a new dish of SNL feeder cells. After this procedure is repeated two to three times, the majority of the dish will consist of undifferentiated colonies. In addition, the qualities of feeder cells, such as density and freshness, are also important.



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Anticipated Results The efficiency of lentiviral transduction to fibroblasts should be >90%. You can estimate the efficiency of infection with the GFPencoding lentivirus. On the other hand, because retroviruses can be transfected only into dividing cells, the transduction efficiency may stay at ∼30% to 60%. From 10 days to 2 weeks after retroviral transduction, some granule colonies usually appear (Fig. 4A.2.2, panel A). However, these colonies are not iPS cells. Generally, clearedged colonies are produced at 3 weeks post transduction (panel B). They can be expanded after being picked up and transferred to another plate. Established iPS cells show hESlike morphologies on feeder cells (panel C). When the cells reach this stage, you should passage once a week. The expression of markers in pluripotent stem cells can be detected in iPS cells as similar level to ES cells. iPS cells typically express SSEA3 (panel D), TRA-1-60 (panel E), TRA-1-81 (panel F), and Nanog (panel G), but not SSEA1 (panel H). Differentiation potentials of iPS cells can be determined easily by embryoid body formation. After a 16-day induction, the expression of differentiation markers such as αfetoprotein (panel I), α-smooth muscle actin (panel J) and βIII-tubulin (panel K) can be confirmed by immunocytochemistry. Another assay for determination of pluripotency, teratoma formation, is also important. Generally, around 3 months after injection of iPS cells into the testes of SCID mice, the mice may appear to be pregnant (panel L). In some cases, black-colored pigment cells can be observed in dissected tumors by the naked eye (panel M). Staining of tumors with hematoxylin and eosin may show that many types of all three germ layers exist in the teratoma if parental iPS cells are pluripotent (panel N). Treatment of human iPS cells with Y-27632, which is an inhibitor for p160-Rhoassociated coiled-coil kinase (ROCK), before harvesting, may improve the survival rate. If you have trouble with frail viability of iPS cells, you can treat the cells at least an hour before harvesting.

Time Considerations


It takes 1 week to successfully transduce the fibroblasts with the lentiviral vector and to verify transduction by microscopic examination or flow cytometry. Then it requires an additional 5 days to prepare the retrovirus vectors

and transduce the fibroblasts. Once plated on SNL feeder cells, it takes ∼3 weeks for iPS cell colonies to appear and additional time for them to grow to a size where they can be passaged. iPS cells are fed every other day and passaged once a week. Overall, it takes over 3 months to establish an iPS cell line.

Acknowledgements We thank Dr. Tetsuya Ishii, Kanon Takeda, and Yuko Shimazu for reading the manuscript, and Tomoko Ichisaka and Noriko Tsubooka and other members of Yamanaka laboratory for valuable discussion and support. Thanks too to Rie Kato, Ryoko Iyama, Eri Nishikawa, Noriyo Maruhashi and the member of CiRA for administrative support. We are also grateful to Drs. Toshio Kitamura for retroviral system, Peter Andrews for antibodies, and Yoshiki Sasai for instruction on teratoma experiments.

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McMahon, A.P. and Bradley, A. 1990. The Wnt-1 (int-1) proto-oncogene is required for development of a large region of the mouse brain. Cell 62:1073-1085. Meissner, A., Wernig, M., and Jaenisch, R. 2007. Direct reprogramming of genetically unmodified fibroblasts into pluripotent stem cells. Nat. Biotech. 25:1177-1181. Morita, S., Kojima, T., and Kitamura, T. 2000. Plat-E: An efficient and stable system for transient packaging of retroviruses. Gene Ther. 7:1063-1066. Nakagawa, M., Koyanagi, M., Tanabe, K., Takahashi, K., Ishisaka, T., Aoi, T., Okita, K., Mochiduki, Y., Takizawa, N., and Yamanaka, S. 2008. Generation of induced pluripotent stem cells without Myc from mouse and human fibroblasts. Nat. Biotech. 26:101-106. Okita, K., Ishisaka, T., and Yamanaka, S. 2007. Generation of germline-competent induced pluripotent stem cells. Nature 448:313-317. Park, I.H., Zhao, R., West, J.A., Yabuuchi, A., Huo, H., Ince, T.A., Leroy, P.H., Lensch, M.W., and Daley, G.O. 2008a. Reprogramming of human somatic cells to pluripotency with defined factors. Nature 451:141-146. Park, I.H., Arora, N., Huo, H., Maheraum, N., Ahfeldt, T., Shimamuki, N., Lensch, M.W., Cowan, C., Hochedinger, K., and Daley, G.O. 2008b. Disease-specific induced pluripotent stem cells. Cell 134:877-886. Takahashi, K. and Yamanaka, S. 2006. Induction of pluripotent stem cells from embryonic and

adult fibroblast cultures by defined factors. Cell 126:663-676. Takahashi, K., Tanabe, K., Ohnuki, M., Narita, T., Tomoda, K., and Yamanaka, S. 2007a. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861-872. Takahashi, K., Okita, K., Nakagawa, M., and Yamanaka, S. 2007b. Induction of pluripotent stem cells from fibroblast cultures. Nat. Protoc. 2:3081-3089. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9. Watanabe, K., Ueno, M., Kamiya, D., Nishiyama, A., Matsumura, M., Wataya, T., Takahashim, J.B., Nishikawa, S., Nishikawa, S., Miguruma, K., and Sasai, Y. 2007. A ROCK inhibitor permits survival of dissociated human embryonic stem cells. Nat. Biotech. 25:681-686. Wernig, M., Meissner, A., Foreman, R., Bambrook, T., Ku, M., Hochedinger, K., Bernstein, R.E., and Jaenisch, R. 2007. In vitro reprogramming of fibroblasts into a pluripotent ES-cell-like state. Nature 448:318-324. Yamanaka, S. 2007. Strategies and new developments in the generation of patient-specific pluripotent stem cells. Cell Stem Cell 1:39-49. Yu, J., Vodyanik, M.A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J.L., Tian, S., Nie, J., Jonsdottir, G.A., Ruotti, V., Stewart, R., Slukvin, I.I., and Thomson, J.A. 2007. Induced pluripotent stem cell lines derived from human somatic cells. Science 318:1917-1920.



Supplement 9

Heterokaryon-Based Reprogramming for Pluripotency

UNIT 4B.1

Carlos Filipe Pereira1 and Amanda G. Fisher1 1

Imperial College School of Medicine, Hammersmith Hospital, London, United Kingdom

ABSTRACT Embryonic stem (ES) cells have the ability to self-renew, execute multiple lineage paths, and dominantly reprogram differentiated cells upon cell fusion. Here, we describe an approach that reprograms human B lymphocytes toward pluripotency by generating inter-species heterokaryons with mouse ES cells. This induces a human ES-specific gene expression profile, in which the extent and the rapidity of conversion allows us to compare the capacity of different mouse ES cell lines to dominantly induce pluripotency. This approach, coupled with pharmacological inhibition, gene knock-out, or knockdown permits factors that are required to directly induce reprogramming to be defined individually, as well as in combination. Experimental heterokaryons provide a simple and tractable approach to address the mechanisms underlying direct reprogramming to pluripotency. The procedure requires 5 days to complete. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 9:4B.1.1-4B.1.14. Keywords: reprogramming r embryonic stem (ES) cell r pluripotency r cell fusion r heterokaryon

INTRODUCTION Epigenetic reprogramming can be achieved in different ways including nuclear transfer or the forced expression of transcription factors to induce pluripotency (Hochedlinger and Jaenisch, 2006; Yamanaka, 2007). However, the low frequency and/or the long period of time required for inducing pluripotency (Stadtfeld et al., 2008) hinders a systematic appraisal of the mechanisms underlying direct reprogramming to pluripotency. Reprogramming can also be achieved by cellular fusion. Spontaneous and experimental cell fusion of differentiated cells with pluripotent cells induces the expression of pluripotencyassociated markers in the hybrid cells (Tada et al., 2001; Cowan et al., 2005; Silva et al., 2006). Here, we describe a simple protocol for generating heterokaryons (cells in which parental nuclei share the same cytoplasm but remain spatially discrete), in which the nuclear events that occur in the donor and recipient nucleus can be discerned using species-specific reagents (Terranova et al., 2006; Pereira et al., 2008). In addition to inducing pluripotency-associated gene expression by somatic cells, this procedure allows an analysis of epigenetic changes within the reprogrammed nucleus over time, including nuclear re-organization, DNA/chromatin modifications, and the requirement of dominant trans-acting factors (Pereira et al., 2008). This unit describes a protocol for the reprogramming of human somatic cells by the formation of heterokaryons with mouse ES cells. The unit begins with the protocol for heterokaryon formation and gene expression analysis by quantitative RT-PCR (Basic Protocol), and it is followed by an alternative method (Alternate Protocol) that allows for the selection of heterokaryons without pre-labeling and fluorescence-activated cell sorting (FACS).

Manipulation of Potency Current Protocols in Stem Cell Biology 4B.1.1-4B.1.14 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc04b01s9 C 2009 John Wiley & Sons, Inc. Copyright

4B.1.1 Supplement 9

BASIC PROTOCOL

GENERATION OF INTER-SPECIES HETEROKARYONS BETWEEN HUMAN B LYMPHOCYTES AND MOUSE ES CELLS This protocol is used for the formation of heterokaryons between human somatic cells and mouse ES cells. Human B cells are fused with mouse ES cells using polyethylene glycol (PEG), and the reprogramming of human somatic cells is monitored by human gene–specific quantitative RT-PCR (Fig. 4B.1.1A). Inter-species heterokaryons can be generated by cell fusion of adherent ES cells and lymphocytes (non-adherent cells). The resulting heterokaryons will attach to gelatin-coated dishes or to irradiated mouse embryonic fibroblast (MEF) feeder layers. NOTE: The quality of donor and recipient cells is essential for reprogramming experiments. We suggest that several ES cell lines/clones that are known to give germ-line transmission (donor) are initially tested in the reprogramming assay. In addition, different somatic cells (recipient) may also be more or less prone to successful reprogramming. NOTE: In addition to gene expression, inter-species heterokaryons can be generated in parallel for other applications (Fig. 4B.1.1A), including bisulfite genomic sequencing (BGS), immunofluorescence (IF; Fig. 4B.1.1C), fluorescence in situ hybridization (FISH), and FACS analysis (Pereira et al., 2008).

Materials

HeterokaryonBased Reprogramming for Pluripotency

Epstein-Barr virus (EBV)–transformed human B cell clones Human B cell medium (hB cell medium; see recipe) Mouse ES/heterokaryon medium (see recipe) Mouse ES cells cultured on gelatin-coated dishes or using mouse embryonic fibroblasts (MEFs) as feeder layers (UNIT 1C.4) Mitotically inactivated MEFs (UNIT 1C.3) 50% (w/v) Polyethylene glycol (PEG) 1500 in 75 mM HEPES, pH 8.0 (Roche, cat. no. 10783641001) Knockout (KO) Dulbecco’s Modified Eagle’s Medium (KO-DMEM; Invitrogen, cat. no. 10829-018) Calcium- and magnesium-free phosphate-buffered saline without (CMF-PBS; Invitrogen, cat. no. 14190-094) 0.05% (w/v) trypsin/EDTA (Invitrogen, cat. no. 25300-054) Vybrant multicolor cell-labeling kit (Molecular Probes, cat. no. V22889) containing: 1,1 -dioctadecyl-3,3,3 ,3 -tetramethylindodicarbocyanine (DiD) cell labeling solution 1,1 -dioctadecyl-3,3,3 ,3 -tetramethylindocarbocyanine perchlorate (DiI) cell labeling solution Leukemia inhibitory factor (LIF; Esgro, Chemicon/Millipore, cat. no. ESG1107) Liquid N2 Mouse monoclonal anti–human LaminA/C (Vector, cat. no. VP-L550), optional Vectashield with DAPI (0.1 μg/ml; Vector, cat. no. H-1200), optional Alexa Fluor 568 phalloidin (Molecular Probes, cat. no. A12380), optional FACS buffer (see recipe) RNA-BEE RNA isolation solvent (AMS Biotechnology, cat. no. CS-501B) DEPC-treated water (Ambion, cat. no. 9915G) TURBO DNA-free kit (Ambion, cat. no. 1907) Superscript first-strand synthesis system (Invitrogen, cat. no. 18080-085) containing: First-strand buffer Superscript III 10 mM dNTP mix (Invitrogen, cat. no. 18427-013)

4B.1.2 Supplement 9


Oligo (dT)12-18 primers (Invitrogen, cat. no. 18418-012) RNaseOUT (Invitrogen, cat. no. 10777-019) SYBR Green Master Mix (Qiagen, cat. no. 204145) Human gene-specific primers (see Table 4B.1.1) 175-cm2 tissue culture flasks Gelatin-coated 90-mm tissue culture dishes (see recipe) 37◦ C water bath 37◦ C, 5% CO2 incubator 10-ml pipet 50- and 15-ml conical tubes (BD Falcon) Hemacytometer Conical 30-ml universal tubes (Sterilin, cat. no. 128A) Pasteur pipets 70-μm cell strainer (BD Falcon, cat. no. 352350) 5-ml polystyrene round-bottom tubes (BD Falcon, cat. no. 352054) FACS DiVa cell sorter (Becton Dickinson) or similar Nanodrop ND-1000 spectrophotometer Dyad DNA engine 96-well plates for PCR (Bio-Rad, cat. no. MLL-9651) Real-time PCR engine (MJ research Chromo4) Additional reagents and equipment for counting cells using a hemacytometer (UNIT 1c.3), growing mouse ES cell culture on feeders (UNIT 1C.4), for mitotically inactive mouse embryonic fibroblasts (UNIT 1C.3), for RNA extraction (Kingston et al., 1996), and for RT-PCR (Giulietti et al., 2001) NOTE: The following procedures (steps 1 to 36) are performed in a Class II biological hazard flow hood or a laminar-flow hood. All solutions and equipment coming into contact with live cells must be sterile, and proper aseptic technique should be used accordingly.

Label mouse ES cells and human B lymphocytes 1. Grow 1 × 108 EBV-hB lymphocytes in suspension using 175-cm2 tissue culture flasks. Plate 2 × 107 EBV-hB cells in 200 ml of hB cell medium. Change hB cell medium every 48 hr by centrifuging 5 min at 200 × g, room temperature. 2. In parallel, grow 1 × 108 mouse ES cells on five 90-mm gelatin-coated dishes or on irradiated MEFs (according to the requirements of the specific ES cell line; UNIT 1C.4). Plate 3 × 106 ES cells per 90-mm dish in 15 ml of ES medium. Change medium the following day and collect cells after 48 hr. We recommend to start with a large number of cells since a good fusion efficiency will be ∼10% (Fig. 4B.1.1B). For a kinetic experiment of three time points, 1 × 108 cells of each cell-type are required.

3. Warm the following in a 37◦ C water bath: PEG solution, serum-free KO-DMEM, complete ES/heterokaryon medium, CMF-PBS, and 0.05% trypsin/EDTA. 4. Collect ES cells by adding 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes and incubate at 37◦ C, 5% CO2 for 5 min. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate trypsin. Dissociate ES cell colonies and disrupt cell clumps to single-cells by pipetting up and down. It is important to get a single-cell suspension of ES cells for cell fusion. Clumps of cells will favor ES × ES cell fusions rather than hB × ES fusions. If required, an additional step of resuspension in trypsin before inactivation with medium may be included.

5. Collect both cell types in 50-ml conical tubes and centrifuge 5 min at 200 × g, room temperature. Current Protocols in Stem Cell Biology


4B.1.3 Supplement 9

A

hB cells DiI+

PEG reprogramming heterokaryon DiI+ DiD+

heterokaryon DiI+ DiD+ mES cells DiD+

time after day 0 cell fusion

day 1

day 2

day 3

FACS sorting of heterokaryons store pellets at -80 C Other applications: IF, FACS, FISH, BGS, etc.

RNA extraction and cDNA synthesis human gene-specific qRT-PCR analysis

C

B human B (hB)

hB + mES

PEG fused (d0) hB:mES 10:1 2.5%

hB:mES 5:1 3.6%

hB:mES 1:1 11.5%

DiD

fluorescence intensity

mouse ES (mES)

DiI

fluorescence intensity

Figure 4B.1.1 (legend at right)


4B.1.4 Supplement 9


Table 4B.1.1 Human Gene-Specific Primers for qRT-PCR Analysisa

Species/Gene hGapdh

NM 002046

hHprt

NM 000194

hOct4 hNanog hCripto hDnmt3b hTert hTle1 hSox2 hRex1 hCD37 hCD19 hCD45

Sequence 5 -3

Accession number

NM 002701 NM 024865 NM 003212 NM 006892 NM 198253 NM 005077 NM 003106 NM 174900 NM 001774 NM 001770 NM 002838

s

TCTGCTCCTCCTGTTCGACA

as

AAAAGCAGCCCTGGTGACC

s

TCCTTGGTCAGGCAGTATAATCC

as

GTCAAGGGCATATCCTACAACAAA

s

TCGAGAACCGAGTGAGAGGC

as

CACACTCGGACCACATCCTTC

s

CCAACATCCTGAACCTCAGCTAC

as

GCCTTCTGCGTCACACCATT

s

AGAAGTGTTCCCTGTGTAAATGCTG

as

CACGAGGTGCTCATCCATCA

s

GTCAAGCTACACACAGGACTTGACAG

as

AGTTCGGACAGCTGGGCTTT

s

GCCAGCATCATCAAACCCC

as

CTGTCAAGGTAGAGACGTGGCTC

s

TGTCTCCCAGCTCGACTGTCT

as

AAGTACTGGCTTCCCCTCCC

s

CACACTGCCCCTCTCACACAT

as

CATTTCCCTCGTTTTTCTTTGAA

s

GCGTACGCAAATTAAAGTCCAGA

as

CAGCATCCTAAACAGCTCGCAGAAT

s

GTGGCTGCACAACAACCTTATTT

as

GCCTAACGGTATCGAGCGAG

s

GCTCAAGACGCTGGAAAGTATTATT

as

GATAAGCCAAAGTCACAGCTGAGA

s

CCCCATGAACGTTACCATTTG

as

GATAGTCTCCATTGTGAAAATAGGCC

a Adapted from Pereira et al., 2008.

Figure 4B.1.1 (at left) Formation of inter-species heterokaryons between human lymphocytes and mouse ES cells. (A) Shows the experimental strategy used for the generation and analysis of inter-species heterokaryons. Human B lymphocytes (hB) and mouse embryonic stem cells (mES) are respectively labeled with the cell membrane dyes DiI and DiD and fused in the presence of polyethylene glycol (PEG). The resulting heterokaryons (cells in which parental nuclei share the same cytoplasm but remain discrete) are cultured under conditions that promote mES selfrenewal and FACS sorted 1, 2, and 3 days after cell fusion. Total RNA is extracted and reverse transcribed for species-specific qRT-PCR analysis. In parallel, heterokaryons may be generated for other applications including immunofluorescence (IF), FACS analysis, fluorescence in situ hybridization (FISH), and bisulfite genomic sequencing (BGS; Pereira et al., 2008). (B) Fusion efficiency is assessed by FACS analysis as the percentage of double-labeled cells (upper-right quadrant of dot plots). Cell fusion with different hB:mES ratios is shown. (C) hB-derived nuclei are distinguished from mouse nuclei by IF on the basis of DAPI (blue) and human Lamin A/C staining (green). A confocal picture of a heterokaryon [one mouse (with DAPI intense foci, blue) and one human nucleus (hLamin A/C positive, green)] 2 days after cell fusion is shown. Actin staining with phalloidin (red) delineates individual cells.



Supplement 9

6. Wash both cell types once in 50 ml CMF-PBS and count cells with a hemacytometer (UNIT 1C.3). 7. Resuspend 1 × 108 of each cell type in 20 ml of prewarmed serum-free KO-DMEM. 8. Respectively label ES cells with 50 μl of Vybrant 1,1 -dioctadecyl-3,3,3 ,3 tetramethylindodicarbocyanine (DiD) and hB lymphocytes with 50 μl of 1,1 dioctadecyl-3,3,3 ,3 -tetramethylindocarbocyanine perchlorate (DiI) cell labeling solutions. Incubate for 15 min in a 37◦ C water bath with occasional mixing. Time and dye concentration required for homogeneous labeling depends on the cell type used and has to be optimized. Cells can be analyzed by flow cytometry using the FL-2 (DiI) and FL-4 (DiD) channels (Fig. 4B.1.1B). In our hands, ES cells and lymphocytes can be homogeneously labeled for 15 min at 37◦ C without causing major cell death.

9. Centrifuge cells 5 min at 200 × g, room temperature, and aspirate supernatant to remove the dye. 10. Resuspend cell pellets in 25 ml of prewarmed complete ES/heterokaryon medium for recovery.

Fuse the cells 11. Centrifuge cells 5 min at 200 × g, room temperature. Remove supernatant and resuspend both cell types in 15 ml of prewarmed CMF-PBS. 12. Mix lymphocytes and ES cells in a conical 30-ml universal tube, ratio 1:1. Varying the cell ratio can have a big impact on cell fusion efficiency (Fig. 4B.1.1B). This is dependent on the physical features of cell types used and has to be optimized. In our hands the optimal lymphocyte:ES ratio is 1:1.

13. Centrifuge cell mixture 5 min at 200 × g, room temperature. With a Pasteur pipet, remove the supernatant completely, including drops on the plastic surface. Complete removal of the supernatant is essential to avoid dilution of PEG.

14. Disrupt the pellet by gently tapping the bottom of the tube. 15. Place the cells in a 37◦ C water bath for 2 min. The optimal temperature for cell fusion is 37◦ C. We recommend that both cell pellet/tube and PEG are prewarmed at 37◦ C for cell fusion.

16. Add 1 ml of PEG dropwise over a 60-sec period. Rotate and rock the tube slowly but continuously during this time to promote cell contact. The quality of PEG and pH of the solution is very important for successful cell fusion. The 50% PEG 1500 solution is commercially available (see Materials section) but also can be manually prepared.

17. Place the tube in a 37◦ C water bath and gently continue mixing the cells in PEG for an additional 90 sec. Exact timing in PEG is vital for the outcome.

18. Add 1 ml of serum-free KO-DMEM (prewarmed at 37◦ C) dropwise to the fusion mixture, rotating and rocking the tube, over a 60-sec period. 19. Add 3 ml of KO-DMEM over a 2-min period, continuously mixing the cells. HeterokaryonBased Reprogramming for Pluripotency

20. Slowly add 10 ml of KO-DMEM. The sequence described above is designed to add KO-DMEM slowly to the mixture. The objective is to dilute PEG but to avoid osmotic shock and cell death.

4B.1.6 Supplement 9


21. Mix by slowly inverting the tube 4 to 6 times. Small clusters of cells induced by PEG treatment can be observed at this point.

22. Incubate 3 min at 37◦ C.

Plate the heterokaryons 23. Centrifuge fusion products 5 min at 250 × g, 37◦ C. 24. With a Pasteur pipet, aspirate the supernatant. Carefully add 10 ml of prewarmed ES/heterokaryon medium plus 1000 U/ml of LIF, without disrupting the pellet. 25. Incubate the pellet of cells in medium 3 min at 37◦ C. 26. Carefully resuspend cell fusions in the required amount of ES/heterokaryon medium plus 1000 U/ml of LIF. Usually, 15 ml per 90-mm dish is sufficient. If required, fusion efficiency can be assessed at this point directly by FACS analysis (Fig. 4B.1.1B).

27. Remove a small aliquot of cells (usually 1/50 of total) and transfer to a 15-ml centrifuge tube. Centrifuge 5 min at 200 × g, room temperature. 28. Wash the aliquot in 15 ml CMF-PBS and aspirate the supernatant. Snap freeze the pellet in liquid N2 (day 0) and store at −80◦ C. 29. Plate cells in 90-mm gelatin-coated dishes at ∼0.5 × 106 cells/cm2 with 15 ml ES/heterokaryon medium plus 1000 U/ml of LIF. Swirl the dish to ensure distribution over the entire plate surface. 30. Incubate dishes overnight at 37 ◦ C, 5% CO2 . Heterokaryons will attach to the dish.

31. Culture heterokaryons in the same conditions used for mouse ES cells. Change medium daily (the majority of unfused lymphocytes will be removed when changing medium). Heterokaryons can be sorted before or after culturing. However, we recommend sorting after culturing as it results in better yield and decreased cell death. Heterokaryons can be monitored over time by fluorescence microscopy combining staining with DAPI, antihuman LaminA/C, and F-actin/phalloidin (Fig. 4B.1.1C). For additional information, refer to Pereira et al. (2008).

Sort heterokaryons 32. At days 1, 2, and/or 3 aspirate medium, wash three times, each time with 10 ml CMF-PBS to remove cell debris and unfused lymphocytes. 33. Add 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes. Incubate 5 min at 37◦ C, 5% CO2 . 34. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate the trypsin. Dissociate cell clumps to single-cells by pipetting up and down. 35. Collect cells in 50-ml conical tubes. Centrifuge 5 min at 200 × g, room temperature. 36. Wash cells with 20 ml CMF-PBS and resuspend the cells in 5 ml of FACS buffer. 37. Pass the cells through a 70-μm cell strainer to remove clumps of cells. Transfer the cells to 5-ml polystyrene round-bottom tubes and put them on ice. 38. Sort the double-labeled population (heterokaryons, Fig. 4B.1.1B) using a FACS DiVa cell sorter (or equivalent) at 4◦ C.



Supplement 9

39. Wash recovered cells in 10 ml ice-cold CMF-PBS by centrifuging 5 min at 200 × g, 4◦ C. 40. Aspirate the supernatant completely and snap freeze the pellet in liquid N2 (day 1, 2, and 3) and store at −80◦ C.

Analyze expression in heterokaryons 41. After collection of the last sample (day 3), extract RNA from frozen samples (day 0, 1, 2, and 3) using RNA-BEE. Refer to Kingston et al. (1996) for a detailed procedure of RNA extraction. 42. Resuspend RNA pellets in 20 μl of DEPC-treated water and eliminate residual contaminating DNA by treatment with TURBO DNase-free kit. 43. Measure the RNA concentration with a Nanodrop spectrophotometer. 44. Reverse transcribe 1 to 3 μg of RNA using the Superscript first-strand synthesis system: dilute RNA in RNase-free water to a final volume of 11 μl and add 1 μl of 10 mM dNTP mix and 1 μl of Oligo (dT)12-18 . 45. Incubate 5 min at 65◦ C and on ice for 1 min. Add 1 μl of 0.1 M DTT, 4 μl of 5× first-strand buffer, 1 μl of RNaseOUT and 1 μl of 200 U/μl Superscript III. 46. Incubate samples 15 min at 25◦ C, 1 hr at 50◦ C,and 15 min at 75◦ C on a Dyad DNA engine. 47. Dilute cDNA 1/10 in water. Samples can be stored at −20◦ C for years.

48. Determine the abundance of human gene-specific transcripts in the heterokaryons at day 0, 1, 2, and 3 using quantitative RT-PCR (for review on real-time quantitative PCR, see Giulietti et al., 2001). We preferentially use SYBR Green Master Mix in a 35-μl reaction [2 μl of cDNA, 1.05 μl of combined primer pairs (10 μM each, Table 4B.1.1), 17.5 μl of Master Mix and 14.45 μl of water], using 96-well plates on a Chromo4 DNA engine under the following cycling conditions: 1 cycle: 40 cycles:

15 min 15 sec 30 sec 30 sec

95◦ C 94◦ C 60◦ C 72◦ C

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

Follow the PCR reaction with a plate reading. Perform each PCR reaction in triplicate. If designing new primer sets, ensure that the selected primers are species-specific and will therefore not cross-anneal with mouse sequences (Table 4B.1.1). Adjust cDNA dilution if the abundance of hGapdh is very different between samples.

49. Acquire data using the MJ Opticon Monitor 3 software. Use the comparative threshold method: for each primer pair set the threshold at the onset of the log-linear phase. The data can be exported to Excel spreadsheets. Calculate the amount of target genes normalized to the endogenous housekeeping gene (hGapdh) using the following equation: 2−C(T) × 1000 HeterokaryonBased Reprogramming for Pluripotency

where C(T) represents the threshold cycle at which fluorescence due to PCR products becomes detectable above background and C(T) is the C(T) of the target gene subtracted by the C(T) of the housekeeping gene (Fig. 4B.1.2).

4B.1.8 Supplement 9



Relative expression (/hGapdh)

0

0.01

0.02

0.03

0.04

0.05

4 3.5 3 2.5 2 1.5 1 0.5 0

16 14 12 10 8 6 4 2 0

d0

d1

d2

hTert

hDnmt3b

ES Oct4

ES WT

hOct4

d3

d0

d3

0.4

0.35 0.3 0.25 0.2 0.15 0.1 0.05 0

0.035 0.03 0.025 0.02 0.015 0.01 0.005 0

d1

d2

hRex1

hSox2

0.8

0.4

d0

d1

hHprt

hTle1

d2

hCripto

Time after fusion (days)

0

5

10

15

20

25

30

0

0.5

1

1.5

2

2.5

0

1.2

0.8

0

1.6

2

1.2

hNanog

1.6

2

d3

0

4

8

12

16

20

6 5 4 3 2 1 0

3.5 3 2.5 2 1.5 1 0.5 0

d0

d1

d2

hCD37

hCD45

hCD19

d3

Figure 4B.1.2 Kinetics of human lymphocyte reprogramming by mouse ES cells. Mouse ES cells expressing Oct4 (black bars), or in which Oct4 expression has been ablated (white bars, negative control; Niwa et al., 2000) are fused to hB lymphocytes and heterokaryons collected over the period of 3 days after cell fusion. The activation of human ES-specific genes (hOct4, hNanog, hCripto, hDnmt3b, hSox2, hTle1, hTert, and hRex1) and silencing of lymphocyte-specific genes (hCD19, hCD45, and hCD37) is quantified by qRT-PCR. Data is normalized to hGapdh and the expression of hHprt is added as a control gene. Error bars indicate the standard deviation (s.d.) of 2 to 3 independent experiments. Adapted from Pereira et al. (2008).


4B.1.9

Supplement 9

ALTERNATE PROTOCOL

GENERATION AND ANALYSIS OF INTER-SPECIES HETEROKARYONS WITHOUT FACS SORTING This protocol is used for the formation of heterokaryons between human somatic cells and mouse ES cells. This simplification of the Basic Protocol allows the selection of heterokaryons in culture based on a combination of drugs, overcoming the requirements for prelabeling and FACS sorting. Using drugs or specific cell lines carrying selective markers, it is possible to generate a population of heterokaryons suitable for gene expression analysis. Here, we give an example—the use of Ara-C in combination with ouabain for heterokaryon selection. This protocol is adequate when large numbers of cells are required for a specific analysis, for systematic cell fusion experiments, or when FACS sorting facilities are not easily accessible. Similar results have been obtained when this protocol was performed in parallel with the Basic Protocol.

Additional Materials (also see Basic Protocol) Ara-C (cytosine β-D-arabinofuranoside, Sigma, cat. no. C-1768) Ouabain (G-Strophanthin; Sigma, cat. no. O-3125) HAT (20 μM hypoxanthine, 0.08 μM aminopterine and 3.2 μM thymidine) media supplement (Sigma, cat. no. H0262-10VL), optional Puromycin (Sigma, cat. no. P9620), optional Generate inter-species heterokaryons without FACS sorting 1. Collect lymphocytes and ES cells following steps 1 to 6 from the Basic Protocol. Using this simplified protocol, cell labeling will not be necessary.

2. Mix cells and proceed with cell fusion. Follow steps 11 to 28 from the Basic Protocol. 3. Plate cells in 90-mm gelatin-coated dishes at ∼0.5 × 106 cells/cm2 with 15 ml ES/heterokaryon medium plus 1000 U/ml of LIF. Swirl the dish to ensure distribution over the entire plate surface. 4. Incubate 6 hr at 37◦ C, 5% CO2 . 5. Add Ara-C (10−5 M; 1 μl of 0.1 M stock per 10 ml of medium) and ouabain (10−5 M; 5.8 μl of 10 mg ml−1 stock per 10 ml of medium) to the dishes. This combination of drugs allows the selection of heterokaryons in culture. Proliferating ES cells are eliminated by Ara-C, a cytosine analog that inhibits DNA synthesis and therefore kills proliferating cells (nondividing cells, i.e., heterokaryons, will not be affected). Ouabain specifically kills human but not mouse cells (or inter-species heterokaryons). As an alternative to Ara-C, Hprt-/- ES cells (Hooper et al., 1987), which die in HAT selective medium, or puromycin-resistant human lymphocytes may be used.

6. Incubate 16 hr at 37◦ C, 5% CO2 . 7. Aspirate medium and wash once with 10 ml CMF-PBS to remove Ara-C. Add 10 ml of ES/heterokaryon medium plus 1000 U/ml LIF and ouabain (10−5 M). Change medium every day. 8. On day 1, 2, and 3, aspirate medium, wash three times, each time with 10 ml CMF-PBS to remove cell debris and unfused lymphocytes. 9. Add 2 ml of 0.05% trypsin/EDTA to 90-mm culture dishes. Incubate 5 min at 37◦ C, 5% CO2 . HeterokaryonBased Reprogramming for Pluripotency

10. Using a 10-ml pipet, add 8 ml of ES/heterokaryon medium to inactivate trypsin. Collect cells in 15-ml conical tubes. Centrifuge 5 min at 200 × g, room temperature. 11. Wash recovered cells in 15 ml CMF-PBS by centrifuging 5 min at 200 × g, room temperature.



12. Aspirate the supernatant completely and snap freeze the pellet in liquid N2 (day 1, 2, and 3) and store at −80 ◦ C. 13. Follow Basic Protocol steps 41 to 49 for RNA extraction, cDNA synthesis, and quantitative RT-PCR analysis.


FACS buffer Add 1.5 ml of fetal bovine serum (FBS) to 48.5 ml calcium- and magnesium-free phosphate-buffered saline (CMF-PBS). Filter the solution using a 0.22-μm filter (Millipore, cat. no. SLGP033RS) and store up to 2 weeks at 4◦ C.

Gelatin, 0.1% (w/v) Incubate 2% (v/v) gelatin (Sigma, cat. no. G1393) for 15 min in a 37◦ C water bath to liquefy the solution. Add 25 ml of 2% (v/v) gelatin to 475 ml of calcium- and magnesium-free phosphate-buffered saline (CMF-PBS). Filter the solution with a bottle-top filter (0.22-μm; Millipore, cat. no. SCGPU05RE) and store up to 4 weeks at room temperature.

Gelatin-coated culture dishes Add enough 0.1% gelatin to cover the bottom of 90-mm dishes. Incubate for at least 20 min at 37◦ C. Before using, aspirate excess gelatin solution. We recommend coating culture dishes right before use.

hB cell medium RPMI-1640 (Invitrogen, cat. no. 31870-025) containing: 10% (v/v) heat-inactivated FBS 2 mM L-glutamine (Invitrogen, cat. no. 25030-123) 50 U/ml−1 penicillin and 50 μg/ml−1 streptomycin (Invitrogen, cat. no. 15140-122) Filter the medium with a 0.22-μm bottle-top filter Store up to 4 weeks at 4◦ C Mouse ES/heterokaryon medium Knockout DMEM (Invitrogen, cat. no. 10829-018) containing: 10% (v/v) fetal bovine serum (FBS) batch tested for ES cell culture 2 mM L-glutamine (Invitrogen, cat. no. 25030-123) 5 ml of non-essential amino acids (Invitrogen) 50 μM 2-mercaptoethanol (Invitrogen, cat. no. 31350-010) 50 U/ml−1 penicillin and 50 μg/ml−1 streptomycin (Invitrogen, cat. no. 15140122) Filter the medium with a 0.22-μm bottle-top filter Store up to 1 week at 4◦ C COMMENTARY Background information Reprogramming somatic cells to become ES-like is an important goal in cell replacement therapy since it affords the opportunity

to generate and use patient-specific ES-derived cells as grafts. Using this strategy, it would be possible to circumvent the problems of immune rejection that are likely to occur unless



Supplement 9

the recipient and donor stem cells are very closely matched. Several experimental reprogramming strategies to convert somatic cells to pluripotency have been outlined including nuclear transfer, cell fusion, and the forced expression of factors (Hochedlinger and Jaenisch, 2006; Yamanaka, 2007). These approaches have been instrumental for dissecting the re-establishment of pluripotency in a somatic cell at the cellular and molecular level. Retroviral-mediated expression of four transcriptional regulators—Oct4, Sox2, c-Myc, and Klf4—was shown to drive mouse and human fibroblasts into an ES-like state, albeit at low frequency (Jaenisch and Young, 2008; Yamanaka, 2007). These studies have illustrated the importance of several factors for reprogramming, but they also suggested that additional ones might be needed for efficient conversion to pluripotency. Reprogramming can also be achieved by cellular fusion, a process that occurs spontaneously in vitro (Ying et al., 2002), in vivo (Weimann et al., 2003), and experimentally using specific agents (Terranova et al., 2006; Pereira et al., 2008). For example, fusion of differentiated cells with pluripotent ES cells induces the expression of pluripotency-associated markers in the hybrid cells and chromatin remodeling at specific sites in the somatic cell genome (Tada et al., 1997; Cowan et al., 2005). To study reprogramming in heterokaryons and hybrid cells it is important to be able to distinguish and verify events that occur in the donor nucleus from those of the recipient. Inter-species heterokaryons formed between human lymphocytes and mouse ES cells therefore provide a tractable approach to study direct reprogramming to pluripotency.

Critical Parameters and Troubleshooting Troubleshooting advice can be found in Table 4B.1.2.

Anticipated Results


A total of 1 × 108 mouse ES cells and 1 × 108 human B lymphocytes are adequate to sort at least 300,000 heterokaryons per time point (day 1, 2, and 3). This allows the parallel gene expression analysis by qRT-PCR of

human pluripotency-associated genes (hOct4, hNanog, hCripto, hDnmt3b, hSox2, hTle1, hTert, and hRex1), lymphocyte-specific genes (hCD19, hCD45, and hCD37), and housekeeping controls (hGapdh, hHprt). As shown in Figure 4B.1.2, fusion of human lymphocytes with wild-type ES cells (black bars) results in the increased expression of human pluripotency-associated genes over this 3-day period. Pluripotency-associated gene activation is mirrored by a reduction in expression of several human lymphocyte-associated genes after heterokaryon formation. As a control, ES cells lacking Oct4 expression [essential transcription factor for ES cell self-renewal and reprogramming ability (Niwa et al., 2000; Pereira et al., 2008)] are fused to human B lymphocytes (white bars) and the activation of human pluripotency-associated genes is impaired. Reprogramming kinetics should be representative of at least two independent experiments.

Time Considerations Expansion of human B lymphocytes and mouse ES cells (steps 1 and 2) takes 7 days. Usually five 90-mm dishes of confluent ES cells and one 175-cm2 flask of lymphocytes are enough to get 1 × 108 cells each cell-type. On day 0, collection of cell types, counting, and cell labeling (steps 3 to 10) will take between 1 and 1.5 hr. The cell fusion procedure and heterokaryon culture (also performed on day 0; steps 11 to 31) will take ∼30 to 45 min. Pause point: cells are left overnight in culture. Days 1 through 3, including reprogramming kinetics and collection of samples (steps 32 to 40) will require 3 days. Cell sorting duration may vary depending on the instrument and cell number, usually 2 to 3 hr per sample. Pause point: cell pellets can be stored at −80◦ C for months. On day 4, RNA extraction, DNase treatment, and reverse transcription (steps 41 to 47) will take 5 hr. Pause point: reverse transcription can be set up in PCR strip tubes and left overnight including a final step at 4◦ C. On day 5, real-time PCR set up, run, and analysis (steps 48 and 49) will require 5 hr. Setting up a real time PCR plate will take ∼45 min and the PCR run takes 3 to 4 hr, which can be left running overnight.



Table 4B.1.2 Troubleshooting Guide to Heterokaryon Formation and Analysis

Problem

Possible cause

Solution

Heterokaryons are not forming

Poor quality of fused cells

Make sure that both ES cells and B lymphocytes are well maintained in culture. Change medium the day before cell fusion.

PEG is being diluted

Dilution of PEG will result in decreased fusion efficiency. Before addition of PEG, remove PBS completely including drops on the surface of tube.

Cell fusion temperature is not optimal

Always warm the cell pellet and PEG in a 37◦ C water bath. Check the water bath temperature. If necessary, add PEG dropwise while keeping the tube in the water bath.

Mechanical dissociation

A period of cell contact is required to complete cell fusion. Handle heterokaryons carefully. Do not mix or pipet heterokaryons vigorously.

Cell labeling is not optimal

Homogeneous labeling with minimum toxicity has to be optimized. Reduce both dye concentration and labeling time.

PEG treatment

Upon PEG treatment a balance between cell death and fusion efficiency has to be found. Reduce time in PEG or predilute PEG in PBS to decrease cell death.

Osmotic shock after PEG treatment

After PEG treatment add DMEM dropwise slowly over a longer period of time (4 ml over 5 min). Make sure to gently rock the tube continuously to slowly dilute PEG.

Cell sorting conditions are not optimal

Check the purity of sorted population in a control experiment by immnunofluorescence. Exclude doublets of cells when sorting heterokaryons.

Low number of heterokaryons sorted

Increase the number of starting cells for cell fusion or optimize cell ratio to increase efficiency.

Poor RNA quality

Check quality of RNA on a gel. If RNA is degraded, skip DNase treatment. Refer to Kingston et al. (1996) for troubleshooting.

Poor quality of ES cell line

Test another mouse ES cell line, or clone, for cell for heterokaryon-based reprogramming.

cDNA is too diluted

The activation of genes will occur gradually and will be detected at late cycles. Use concentrated cDNA to start with and dilute according hGapdh levels.

Refractory human somatic cell line

Some cell types can be more or less prone to reprogramming. Use another cell type or cell line. We have successfully reprogrammed several human EBV–transformed B cell clones.

Sample contamination with unfused hB lymphocytes

Add ouabain to the heterokaryon medium after cell fusion. Wash dishes very well with PBS to remove lymphocytes. Decrease size of gate when sorting cells to avoid the collection of DiI+ DiD− cells.

Excessive cell death

hGapdh levels very low in heterokaryons

Pluripotency-associated genes are not detected

Lymphocyte-associated genes are not silenced



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Literature Cited Cowan, C.A., Atienza, J., Melton, D.A., and Eggan, K. 2005. Nuclear reprogramming of somatic cells after fusion with human embryonic stem cells. Science 309:1369-1373. Giulietti, A., Overbergh, L., Valckx, D., Decallonne, B., Bouillon, R., and Mathieu, C. 2001. An overview of real-time quantitative PCR: Applications to quantify cytokine gene expression. Methods 25:386-401. Hochedlinger, K. and Jaenisch, R. 2006. Nuclear reprogramming and pluripotency. Nature 441:1061-1067. Hooper, M., Hardy, K., Handyside, A., Hunter, S., and Monk, M. 1987. HPRT-deficient (LeschNyhan) mouse embryos derived from germline colonization by cultured cells. Nature 326:292295. Jaenisch, R. and Young, R. 2008. Stem cells, the molecular circuitry of pluripotency and nuclear reprogramming. Cell 132:567-582. Kingston, R.E., Chomczynski, P., and Sacchi, N. 1996. Guanidine methods for total RNA preparation. Curr. Protoc. Mol. Biol. 36:4.2.1-4.2.9. Niwa, H., Miyazaki, J., and Smith, A.G. 2000. Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat. Genet. 24:372-376. Pereira, C.F., Terranova, R., Ryan, N.K., Santos, J., Morris, K.J., Cui, W., Merkenschlager, M., and Fisher, A.G. 2008. Heterokaryon-based reprogramming of human B lymphocytes for pluripotency requires Oct4 but not Sox2. PLoS Genet. 4:e1000170.

Silva, J., Chambers, I., Pollard, S., and Smith, A. 2006. Nanog promotes transfer of pluripotency after cell fusion. Nature 441:997-1001. Stadtfeld, M., Maherali, N., Breault, D.T., and Hochedlinger, K. 2008. Defining molecular cornerstones during fibroblast to iPS cell reprogramming in mouse. Cell Stem Cell 2:230240. Tada, M., Tada, T., Lefebvre, L., Barton, S.C., and Surani, M.A. 1997. Embryonic germ cells induce epigenetic reprogramming of somatic nucleus in hybrid cells. EMBO J. 16:65106520. Tada, M., Takahama, Y., Abe, K., Nakatsuji, N., and Tada, T. 2001. Nuclear reprogramming of somatic cells by in vitro hybridization with ES cells. Curr. Biol. 11:1553-1558. Terranova, R., Pereira, C.F., Du Roure, C., Merkenschlager, M., and Fisher, A.G. 2006. Acquisition and extinction of gene expression programs are separable events in heterokaryon reprogramming. J. Cell Sci. 119:2065-2072. Weimann, J.M., Johansson, C.B., Trejo, A., and Blau, H.M. 2003. Stable reprogrammed heterokaryons form spontaneously in Purkinje neurons after bone marrow transplant. Nat. Cell Biol. 5:959-966. Yamanaka, S. 2007. Strategies and new developments in the generation of patient-specific pluripotent stem cells. Cell Stem Cell 1:39-49. Ying, Q.L., Nichols, J., Evans, E.P., and Smith, A.G. 2002. Changing potency by spontaneous fusion. Nature 416:545-548.




Imaging Neural Stem Cell Fate in Mouse Model of Glioma

UNIT 5A.1

Khalid Shah1 1

Massachusetts General Hospital, Harvard Medical School, Charlestown, Massachusetts

ABSTRACT This unit describes a protocol for following the fate of stem cells in real time in a mouse model of glioma. Stem cells and tumor cells can be transduced with lentiviral vectors bearing two different luciferases, firefly luciferase (Fluc) and Renilla (Rluc) luciferase, respectively. With the cells labeled in this manner, bioluminescence imaging can be used to study the fate of stem cells in glioma-bearing brains in vivo. Curr. Protoc. Stem Cell C 2009 by John Wiley & Sons, Inc. Biol. 8:5A.1.1-5A.1.11. Keywords: neural stem cell r bi-modal vector r luciferase r fluorescent proteins r glioma r in vivo imaging

INTRODUCTION Several studies have demonstrated the effectiveness of neural stem cell (NSC) transplantation in the treatment of neurodegenerative diseases, including spinal cord injury and brain tumors (Snyder and Macklis, 1995; Ehtesham et al., 2002; Lindvall et al., 2004; Hofstetter et al., 2005; Iwanami et al., 2005; Shah et al., 2005). This unit describes a protocol for simultaneously imaging the fate of engineered NSC and glioma cells in a mouse glioma model. NSC and glioma cells transduced with lentiviral vectors bearing different combinations of fluorescent and bioluminescent proteins can be grown as monolayers and maintained over several passages. The unit begins with a method for transducing NSC and glioma cells with bimodal lentiviral vectors for stable expression of these fluorescent and bioluminescent markers in vitro, followed by transplantation of fluorescent and bioluminescent glioma cells and NSC in mice, and, finally, sequential bioluminescent imaging of NSC fate and glioma progression in mice. The integration of different combinations of bioluminescent and fluorescent proteins into NSC and glioma cells makes it possible to distinguish different populations of cells after intracranial transplantation. Lentiviral vector transduction of cells is followed by cell sorting, which is necessary to obtain a pure population of different fluorescent cell types. The protocol details viral transduction, surgical preparation, craniotomy, cell implantation, animal recovery, and imaging procedures to study stem cell kinetics and migration to malignant brain tumors. NOTE: All solutions and equipment coming into contact with live cells must be sterile. NOTE: All culture incubations should be performed in a humidified 37◦ C, 5%, CO2 incubator unless otherwise specified. NOTE: Viral transductions on human stem cells and glioma cells and cell culture procedures are performed in a biosafety level (BL)-2 facility in a laminar-flow hood.

ENGINEERING STEM CELL AND GLIOMA LINES This protocol is used for transducing human NSC and human glioma cells with lentiviral vectors bearing bioluminescent and fluorescent markers for stable expression of these markers in vitro and in vivo. Both cell types are transduced with lentiviral vectors bearing unique combinations of fluorescent and bioluminescent markers, and cells are sorted by cell sorter. Current Protocols in Stem Cell Biology 5A.1.1-5A.1.11 Published online March 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a01s8 C 2009 John Wiley & Sons, Inc. Copyright

BASIC PROTOCOL 1

Genetic Manipulation of Stem Cells

5A.1.1 Supplement 8

Materials Human neural stem cells (NSC; Rubio et al., 2000) NSC culture medium (see recipe) 0.25% (w/v) trypsin/EDTA (Invitrogen) Human glioma cells (Gli36; Shah et al., 2004) Glioma cell culture medium: DMEM containing 10% FBS and 1× penicillin/streptomycin Plasmid for NSC cells: lentiviral plasmid bearing a fusion between GFP and Fluc (GFP-Fluc; Shah et al., 2008) Plasmid for glioma cells: lentiviral plasmid bearing a fusion between Rluc and DsRed2 (Rluc-DsRed2; Shah et al., 2008) Phosphate-buffered saline (PBS; e.g., Invitrogen) NSC culture medium (see recipe) containing 8 μg/ml polybrene (add from 8 mg/ml polybrene stock in PBS; Fisher) Glioma cell culture medium (see above) containing 8 μg/ml polybrene (add from 8 mg/ml polybrene stock in PBS; Fisher) 5-cm culture dishes (Corning) Fluorescence microscope with appropriate filters for GFP and rhodamine Cell sorter (e.g., FACScalibur from BD Biosciences) Additional reagents and equipment for fluorescence-activated cell sorting (Robinson et al., 2009) Culture cells and lines 1a. For human fetal neural stem cell line: Culture human fetal neural stem cell line (NSC), derived from the human diencephalic and telencephalic regions of 10 to 10.5 weeks gestational age from an aborted human Caucasian embryo, in NSC culture medium in a 5-cm culture dish at 37◦ C in a humidified incubator, to 70% to 80% confluency. These cells are grown as monolayers and are passaged every 4 days by trypsinizing cells in 0.25% trypsin/EDTA. Cells are centrifuged 10 min at 300 × g and plated at 20% density in NSC culture medium. The in vitro and in vivo properties of NSC (including the absence of transformation, clonality, multipotency, stability, and survival) have been described in detail elsewhere (Rubio et al., 2000; Villa et al., 2004; Navarro-Galve et al., 2005).

1b. For human glioma cell line: Culture Gli36, a human glioma cell line whose in vitro and in vivo characteristics have been described elsewhere (Shah et al., 2004, 2005) in glioma cell culture medium at 37◦ C to 70% to 80% confluency. Glioma cells grow as monolayers and are passaged every 4 days by trypsinizing cells in 0.25% trypsin/EDTA. Cells are seeded at 20% density in glioma cell culture medium.

2. When cells reach 70% to 80% confluency, subculture cells at a 1:4 (NSC) or 1:5 (glioma cell) ratio.

Prepare lentiviral vectors 3. Use the CS-CGW transfer plasmid–based lentiviral vector system (Miyoshi et al., 1998) to create lentiviral transfer vectors bearing fusions between Renilla luciferase (Rluc) and Discosoma Red (DsRed2) proteins (LV-Rluc-DsRed2) and lentiviral vectors bearing fusions between firefly luciferase (Fluc) and green fluorescent protein (GFP; LV-GFP-Fluc). Imaging Neural Stem Cell Fate in Mouse Model of Glioma

The construction of LV-GFP-Fluc and LV-Rluc-DsRed2 is described in detail elsewhere (Shah et al., 2008).

5A.1.2 Supplement 8


4. Amplify the cDNA sequences encoding GFP-Fluc fusion and Rluc-DsRed2 fusion by PCR (Kramer and Coen, 2001) and ligate in-frame into NheI/XhoI-digested CS-CGW plasmid (Shah et al., 2008). 5. Produce lentiviral vectors (Shah et al., 2008). 6. Titer the viruses (Shah et al., 2008) and store in PBS at −80◦ C.

Perform viral transduction and cell sorting For NSC 7a. Plate NSC at 60% confluency in a 5-cm dish. 8a. At a time point 18 hrs later, transduce NSC with LV-GFP-Fluc (at MOI = 1) in NSC culture medium containing 8 μg/ml polybrene. 9a. Confirm viral transduction by visualizing cells for GFP expression by fluorescence microscopy 36 to 48 hr after transduction. 10a. At 72 hr after transduction, perform single-cell sorting based on GFP fluorescence, using a cell sorter (also see Robinson et al., 2009) to obtain a monoclonal cell populations. Culture sorted cells in NSC culture medium. We use BD FACScalibur cell sorter (BD Biosciences).

For glioma cells 7b. Plate glioma cells at 40% confluency in a 5-cm dish. 8b. 18 hr later, transduce glioma cells with LV-Rluc-DsRed2 (at MOI = 1) in glioma cell culture medium containing 8 μg/ml polybrene. 9b. Confirm viral transduction by visualizing cells for DsRed2 expression by fluorescence microscopy 48 hr after transduction. 10b. At 72 hr after transduction, trypsinize cells using trypsin/EDTA, wash the cells twice with PBS (each time centrifuging 10 min at 300 × g), and perform single-cell sorting based on rhodamine fluorescence using a cell sorter (also see Robinson et al., 2009) to obtain monoclonal cell populations. Culture sorted cells.

BIOLUMINESCENCE IMAGING IN CULTURE This protocol is used for bioluminescence imaging of NSC and glioma cells expressing different combinations of bioluminescent and fluorescent markers in vitro (see Fig. 5A.1.1).

SUPPORT PROTOCOL

Materials NSC and glioma cells bearing bioluminescent and fluorescent markers (Basic Protocol 1) NSC culture medium (see recipe) 150 mg/ml D-luciferin stock (firefly luciferase substrate; Biotium, cat. no. 10110-1; http://www.biotium.com) in PBS Glioma cell culture medium: DMEM containing 10% FBS 1 mg/ml coelenterazine stock (substrate for Renilla luciferases; Biotium, cat. no. 10102-2; http://www.biotium.com) in ethanol 48- or 96-well clear-bottom black-walled plate Bioluminescence imaging system with IVIS-200 or IVIS-100 (Caliper; http://www.caliperls.com/) or similar bioluminescence imaging system Genetic Manipulation of Stem Cells


Supplement 8

Figure 5A.1.1 Fluorescence and bioluminescence characteristics of NSC (A) and glioma cells (B) in culture. NSC and glioma cells were transduced in culture with LV-GFP-Fluc and Lv-RlucDsRed2, respectively, at MOI = 1, and visualized for GFP (A) or DsRed2 (B) ßuorescence. (C,D) Different concentrations of NSC expressing GFP-Fluc (1.0–1.5 × 105 ) and glioma cells (1.5–6 × 105 ) expressing Rluc-DsRed2 were plated, and, 12 hr later, cells were incubated with 150 μg/ml D-luciferin or 1 μg/ml of coelenterazine and imaged under the CCD with a scan time of 1 min. MagniÞcation, 20×. Adapted from Shah et al. (2008), with permission from Society for Neuroscience.

To image the bioluminescence of transduced NSC 1a. Using a black-walled, clear-bottom 96-well tissue culture plate, seed NSC at several densities spanning 1000 to 10,000 cells per well in 100 μl NSC culture medium, to determine the correlation between the number of transduced NSC and the firefly luciferase bioluminescence signal. Incubate. 2a. At a time point 18 to 24 hr later add D-luciferin (substrate for firefly luciferase) to the culture medium at a 1/10 volume, for a final concentration of 0.15 mg/ml, using a multichannel pipettor. Dilute from a 150 mg/ml D-luciferin stock.

3a. Rock the plate and take images in bioluminescence imager with the appropriate exposure. For firefly luciferase, peak light production from intact cells occurs ∼10 min after substrate addition.

To image bioluminescence imaging of transduced glioma cells 1b. Using a black-walled, clear-bottom 96-well tissue culture plate, seed glioma cells at several densities ranging from 1000 to 10,000 cells per well in 100 μl glioma cell culture medium to determine the correlation between number of transduced glioma cells and Renilla luciferase bioluminescence signal. Incubate. Imaging Neural Stem Cell Fate in Mouse Model of Glioma

2b. At a time point 18 to 24 hr later, add coelenterazine (substrate for Renilla luciferase) to the culture medium at a 1/10 volume, for a final concentration of 0.1 μg/ml, using a multichannel pipettor.

5A.1.4 Supplement 8


The 1 mg/ml ethanol stock of coelenterazine is first diluted to an appropriate concentration with PBS before being added to the culture medium.

3b. Rock the plate and take images in bioluminescence imager with the appropriate exposure. Peak bioluminescence from Renilla luciferases occurs rapidly within the first minute after adding coelenterazine.

CELL TRANSPLANTATION AND IMAGING This protocol is used for transplantation and subsequent imaging of NSC and glioma cells expressing different combinations of bioluminescent and fluorescent markers in mice. It also describes the dual imaging of NSC fate and glioma progression in the mouse glioma model.

BASIC PROTOCOL 2

NOTE: All protocols involving live animals must be reviewed and approved by an Institutional Committee for Ethical Animal Care and Use (IACUC) and must conform to government regulations for the care and use of laboratory animals. NOTE: Mouse surgical procedures are performed in a surgical room designated for animal surgeries. Proper aseptic techniques should be used accordingly.

Materials SCID mice (6-to 8-weeks-old; Charles River Laboratories) Anesthetics: ketamine and xylazine (also see Donovan and Brown, 1998) Betadine solution (Bruce Medical; http://www.brucemedical.com/) 70% isopropyl alcohol (Fisher) Phosphate-buffered saline (PBS), sterile Gli36-Rluc-DsRed2 glioma cells (Basic Protocol 1) Bone wax (Ethicon) Coelenterazine (100 μg/animal in 150 μl saline; Biotium, cat. no. 10102-2) D-luciferin (150 μg/g body weight in 150 μl saline; Biotium, cat. no. 10110-1) Animal shaver Stereotaxic frame (Harvard Apparatus, cat. no. 726049) Stereo dissecting microscope: variable magnification (1 to 4.5; Nikon) Fine scissors (Fine Science Tools, cat. no. 14084-08) Forceps, angled and straight and ultrafine angled (Fine Science Tools) Cotton-tipped applicators Hand-held micro-drill (Fine Science Tools, cat. no. 18000-17) with 0.45-mm round drill burr (VWR) 10-μl Hamilton gastight 1701 syringe with 26-G needle 4–0 vicryl sutures or surgical staples Bioluminescence imaging system with IVIS-200 or IVIS-100 (Caliper; http://www.caliperls.com/) or similar bioluminescence imaging system Additional reagents and equipment for anesthesia of mice (Donovan and Brown, 1998) Anesthetize the animal 1. Grasp the animal firmly with one hand and anesthetize by injecting ketamine and xylazine intraperitoneally (Donovan and Brown, 1998). The ideal dosage for each animal will vary primarily based upon the animal’s body mass (120 mg/kg ketamine and 16 mg/kg xylazine) and age.



Supplement 8

Figure 5A.1.2

Anesthetized mouse in a stereotaxic device being implanted with glioma cells.

2. Use the toe-pinch method to assess the level of sedation by firmly applying pressure to the animal’s toe pads and observing whether or not there is a demonstration of a pain response by the animal. Also monitor breathing and posture. 3. Secure animal on a stereotactic head frame placed under a stereo dissecting microscope and shave dorsal surface of the animal’s head (see Fig. 5A.1.2). 4. Disinfect the shaved area by applying two alternating coatings with Betadine and 70% isopropyl alcohol. 5. Using scissors and forceps, remove the skin from the disinfected region and use a dry cotton swab to completely remove the periosteum membrane from the exposed skull surface. Keep the skull moist by frequent application of sterile PBS following the removal of the periosteum. 6. For glioma cell implantations, use a handheld micro-drill to drill through the bone at the location of the proposed implantation site until the cortical surface is exposed.

Implant tumor cells 7. Place 4 to 5 μl of Gli36-Rluc-DsRed2 glioma cells (100,000 cells) in a 10-μl 26-G Hamilton Gastight 1701 syringe and insert the needle to a specified depth into the left frontal lobe. In our experiments we have used the following stereotactic coordinates: 2.5 mm lateral and 0.5 mm caudal to bregma; depth 2.5 mm from dura.

8. Implant cells over a period of 4 min with 30-sec intervals. Care should be taken to consistently implant tumors at the same location and depth to facilitate bioluminescence interpretation from within this relative point source.

9. After implantation is complete, wait for 5 min and remove needle over a period of 10 min with intervals of 1 min. 10. Seal the burrow hole with bone wax and close the wound with 4–0 vicryl sutures or surgical staples. Imaging Neural Stem Cell Fate in Mouse Model of Glioma

5A.1.6 Supplement 8


Figure 5A.1.3 NSC migrate into gliomas in vivo. Bioluminescence imaging of mice implanted with GFP-Fluc expressing NSC in mice with established Rluc-DsRed2 gliomas. Fluc images of mice on day 3 (A), day 7 (B), and day 10 (C), and Rluc image on day 10 (D). Adapted from Shah et al. (2008) with permission from Society for Neuroscience.

Image in vivo tumor cell bioluminescence Imaging can be performed 24 hr after cell implantation. 11. Anesthetize mouse by injecting the appropriate dose of ketamine and xylazine intraperitoneally (see step 1). 12. Use the toe-pinch method to assess the level of sedation by firmly applying pressure to the animal’s toe pads and observing whether or not there is a demonstration of a pain response by the animal. Also monitor breathing and posture. It is slightly more difficult, yet equally important, to monitor anesthesia during imaging as during surgery. During extended time-course sessions, imaging may be jeopardized by a possible toe-pinch reaction and it may be more appropriate to monitor the animal’s breathing and posture.

13. First, acquire a surface image of each animal using dim polychromatic illumination. Next, measure the spatial distribution of luciferase activity within the mouse brain by photon count recording using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3), according to the manufacturer’s instructions. 14. Image mice for Rluc activity by injecting 100 μg coelenterazine (in 150 μl saline) intravenously via the tail vein and record photon counts 5 min later over a 5-min period using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3) according to the manufacturer’s instructions.

Implant stem cells 15. Anesthetize the same animals implanted with glioma cells with the appropriate dose of ketamine and xylazine (see step 1). 16. Secure on a stereotactic head frame placed under a stereo dissecting microscope. 17. Using a handheld micro-drill, drill hole in the contralateral, right frontal lobe at the following coordinates: 2.5 mm lateral and 0.5 mm caudal to bregma; depth 2.5 mm from dura. Depending on the migrating ability and speed of migration, NSC can be placed at any distance from the gliomas in order to assess migration of NSC to gliomas in the brain. In our studies, we have placed the NSC in the contralateral right frontal lobe of the glioma-bearing mice in order to follow migration of NSC toward gliomas.



Supplement 8

18. Place 4 to 5 μl of NSC expressing GFP-Fluc (NSC-GFP-Fluc; 500,000 cells) in a 10-μl 26-G Hamilton Gastight 1701 syringe and implant cells over 4 min with 30-sec intervals. 19. Wait 5 min and withdraw the syringe over 10 min, with 1-min intervals.

Image in vivo stem cell bioluminescence 20. To image mice for firefly luciferase (Fluc) activity, inject the mice intraperitoneally with 150 μg/g body weight D-luciferin (in 150 μl saline). 21. Acquire images 10 min after D-luciferin administration over a period of 5 min. 22. Measure the spatial distribution of luciferase activity within the brain of the animal by recording photon counts using IVIS-200 or IVIS-100 or similar bioluminescence imaging system (see Fig. 5A.1.3), according to the manufacturer’s instructions. Mice can be imaged every day for Fluc and Rluc activity. Typical exposure times vary between 1 and 10 min. If imaging for both, screen for Rluc activity and then Fluc activity. Allow a 24-hr period between imaging sessions to make sure there is no residual luciferase activity from the previous session.

Allow the animal to recover 23. Observe the animal for recovery. Make certain the animal is restrained and that it cannot cause harm to itself. When the animal is maintaining its own normal body temperature and has a reflexive response to toe-pinch stimulation, return it to a clean and unoccupied cage. For the most part, the animal should survive the procedure despite the absence of an external heat source. The usual recovery time for this procedure can range from 2 to 12 hr. If the animal has not resumed normal grooming and eating behavior beyond this time frame, it may require additional medical attention or euthanasia.

Analyze data 24. Use the software accompanying the imaging equipment to perform the region of interest (ROI) analysis. In our studies, following data acquisition, post-processing and visualization is performed using a home-written program with image display and analysis suite developed in IDL (Research Systems Inc.). Regions of interest are defined using an automatic intensity contour procedure to identify bioluminescence signals with intensities significantly greater than the background. The mean, standard deviation, and sum of the photon counts in these regions are then calculated. For visualization purposes, the bioluminescence images are fused with the corresponding white-light surface images as a transparent pseudocolor overlay, permitting correlation of areas of bioluminescence activity with anatomy. Maintaining a standard region of interest within an experiment (or series of experiments) is important to facilitate comparison of mouse imaging data.


NSC culture medium


DMEM/F-12 (Invitrogen) supplemented with: 0.6% (w/v) D-glucose (Sigma-Aldrich) 0.5% (w/v) AlbuMax (Life Technologies) 0.5% (w/v) L-glutamine (Life Technologies) 20 ng/ml recombinant human FGF (R & D Systems) continued

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20 ng/ml recombinant human EGF (R & D Systems) 1× N2 supplement (Invitrogen) 1% (w/v) nonessential amino acids (Cellgro) 1 mM sodium pyruvate (Cellgro) 26 mM sodium bicarbonate) COMMENTARY Background Information Neural stem cells (NSC) are defined by their ability to self-renew and give rise to mature progenitors of neural lineages. The ability of NSC to migrate to diseased areas of the brain has been documented (Snyder and Macklis, 1995: Aboody et al., 2000; Tang et al., 2003; Shah et al., 2005). Their capacity to differentiate into all neural and glial phenotypes (Gage, 2000) provides a powerful tool for targeting the treatment of both diffuse and localized neurologic disorders. Several studies have demonstrated the effectiveness of NSC transplantation in the treatment of neurodegenerative diseases, including spinal cord injury and brain tumors (Snyder and Macklis, 1995; Ehtesham et al., 2002; Lindvail et al., 2004; Hofstetter et al., 2005; Iwanami et al., 2005; Shah et al., 2005). Taking advantage of their homing properties, NSC have also been modified to deliver selective anti-neoplastic proteins (Ehtesham et al., 2002; Shah et al., 2005), although with mixed results. While these studies demonstrate the feasibility of NSC-based therapy, cellular delivery of therapeutic proteins via NSC grafts will likely require long-term transgene expression. In vivo assays, which permit rapid assessment of the fate of transplanted stem cells, will be useful in designing future stem-cell-based therapies. Bioluminescence imaging exploits the emission of visible photons at specific wavelengths based on energy-dependent reactions catalyzed by luciferases. It is a powerful method for detecting and quantifying the spatial and temporal occurrence of cellular and molecular events and can be efficiently used for longitudinal comparison of cell survival and migration. Luciferases from Renilla (Rluc) and firefly (Fluc) have different substrates, coelenterazine and D-luciferin, respectively, and can be imaged in tumors in the same living mouse with kinetics of light production being separable by timed injections of these two substrates (Shah et al., 2005). This dual-imaging approach has direct applications in studying gene expression from vectors and simultaneously monitoring therapeutic effects in vivo.

Critical Parameters and Troubleshooting An efficient and robust way to follow cells both in culture and in vivo is to transduce them with lentiviral vectors expressing fusions of bioluminescent and fluorescent marker genes. These vectors have the ability to integrate transgenes into the genome of dividing and nondividing cells (Naldini et al., 1996) and provide means of efficient long-term expression in cells and their progeny without using any antibiotic selection marker. The fluorescent marker serves to determine the efficiency of transduction, and, in conjunction with the bioluminescent marker, serves as an in vivo cell-tracking protein. Furthermore, the expression of fluorescent markers in different cell populations also aids in performing pathological analysis on tissue sections in sacrificed animals. Knowing the depth and optical properties of the tissue through which the light will pass is essential in calculating numbers of cells needed to obtain a detectable signal. Generally, firefly luciferase light will be attenuated approximately 10-fold for each centimeter of tissue, but optically dense tissues such as liver will attenuate light much more than skin, bone, or lung. Thus, the number of luciferaseexpressing cells and their localization within the body is critical in obtaining a detectable signal to follow fate of cells in vivo; the deeper the tumors are within the body, or the deeper the intracranial tumor, the greater the signal attenuation. For example, in subcutaneous tumors, cell numbers as low as 1000 firefly luciferase–expressing cells can be detected. Also, D-luciferin has more favorable biodistribution than coelenterazine, and an intraperitoneal injection of luciferin is much more reproducible than the tail vein injection that is required for delivering coelenterazine. Transplanting cells expressing Fluc and Rluc in various sites, using various gene delivery vectors and transgenic models, demonstrates the high accessibility of D-luciferin (Fluc substrate) to various tissues, including the brain. On the other hand, coelenterazine



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is also accessible to many tissues because of its diffusable nature (Lorenz et al., 1996), but its distribution in the intact brain is limited by drug-transport proteins, which can hinder in vivo imaging of Renilla luciferase (Pichler et al., 2004). This problem may be overcome by injecting mice with a blood-brain barrier (BBB) disrupter, e.g., mannitol. Also, mouse fur attenuates and scatters light, and this effect is most pronounced in black mice. This problem may be overcome by using nude mice or shaving animals over the region(s) of interest for imaging. Luciferase imaging in mice offers the possibility of imaging mice serially. To perform repetitive imaging of mice, the user should take into account that luciferase levels in mice peak ∼10 min after intraperitoneal. injection, then decline slowly to background levels by 6 to 8 hr post injection (Paroo et al., 2004). Coelenterazine has a more rapid kinetic course in mice. Therefore, maximum imaging signal for Renilla luciferases is obtained immediately after injecting coelenterazine through intravenous or intra-cardiac routes (Bhaumik and Gambhir, 2002). For imaging two different molecular events simultaneously, for example, stem cell fate and glioma volumes in the same mouse, it is advisable to image Renilla luciferase activity first, and then image firefly luciferase activity. Bioluminescence signal from mice implanted with NSC-expressing bioluminescent proteins in the brain varies with the presence and absence of tumors, and in different mice. We have previously shown that human neural stem cell survival is much improved in SCID mice as compared to nude mice (Shah et al., 2008), which could be attributed to the fact that SCID mice (lacking functional T and B cells) are more immune-compromised than nude mice (which lack functional T cells only), and this may implicate immune rejection as a factor in NSC survival in the brain. Our studies also reveal the persistence of NSC in the brains of tumor-bearing mice as compared to normal mice, implying that glioma cells or host response may modulate human stem cell survival either through secretion of growth factors or by inhibition of molecules involved in foreign cell rejection. While designing experiments for imaging human stem cell fate in mouse tumor models, the choice of mouse and the tumor cell type should be taken into consideration.

Anticipated Results The protocols in this unit generate useful information on the fate of NSC in a mouse model of glioma, and are suitable for a number of other disease models. Both stem cells and glioma cells can be easily transduced with lentiviral vectors, and bioluminescence imaging can be used to study the fate of stem cells in different disease models in vivo. Glioma cell survival is higher than NSC survival in mice. Furthermore, we have shown that the presence of glioma cells improves the survival of NSC in the brain.

Time Considerations It takes 1 week for glioma cells and 2 weeks for NSC to grow before they are transduced with lentiviral vectors. Glioma cells are implanted 2 to 3 days after lentiviral transduction, and transduced NSC are implanted 3 to 4 days after glioma cell implantation. Both glioma cells and NSC can be followed in real time in vivo for a period of 3 to 4 weeks before glioma growth results in the mortality of animals.

Literature Cited Aboody, K.S., Brown, A., Rainov, N.G., Bower, K.A., Liu, S., Yang, W., Small, J.E., Herrlinger, U., Ourednik, V., Black, P.M., Breakefield, X.O., and Snyder, E.Y. 2000. Neural stem cells display extensive tropism for pathology in adult brain: Evidence from intracranial gliomas. Proc. Natl. Acad. Sci. U.S.A. 97:12846-12851. Bhaumik, S. and Gambhir, S.S.. 2002. Optical imaging of Renilla luciferase reporter gene expression in living mice. Proc. Natl. Acad. Sci. U.S.A. 99:377-382. Donovan, J. and Brown, P. 1998. Anesthesia. Curr. Protoc. Immunol. 27:1.4.1-1.4.5. Ehtesham, M., Kabos, P., Gutierrez, M.A., Chung, N.H., Griffith, T.S., Black, K.L., and Yu, J.S. 2002. Induction of glioblastoma apoptosis using neural stem cell-mediated delivery of tumor necrosis factor-related apoptosis-inducing ligand. Cancer Res. 62:7170-7174. Gage, F.H. 2000. Mammalian neural stem cells. Science 287:1433-1438. Hofstetter, C.P., Holmstrom, N.A., Lilja, J.A., Schweinhardt, P., Hao, J., Spenger, C., Wiesenfeld-Hallin, Z., Kurpad, S.N., Frisen, J., and Olson, L. 2005. Allodynia limits the usefulness of intraspinal neural stem cell grafts: Directed differentiation improves outcome. Nat. Neurosci. 8:346-353. Iwanami, A., Kaneko, S., Nakamura, M., Kanemura, Y., Mori, H., Kobayashi, S., Yamasaki, M., Momoshima, S., Ishii, H., Ando, K., Tanioka, Y., Tamaoki, N., Nomura, T., Toyama, Y., and Okano, H. 2005. Transplantation of human



neural stem cells for spinal cord injury in primates. J. Neurosci. Res. 80:182-190. Kramer, M.F. and Coen, D.M. 2001. Enzymatic amplification of DNA by PCR: Standard procedures and optimization. Curr. Protoc. Mol. Biol. 56:15.1.1-15.1.14. Lindvall, O., Kokaia, Z., and Martinez-Serrano, A. 2004. Stem cell therapy for human neurodegenerative disorders: How to make it work. Nat. Med. 10:S42-S50. Lorenz, W.W., Cormier, M.J., O’Kane, D.J., Hua, D., Escher, A.A., and Szalay, A.A. 1996. Expression of the Renilla reniformis luciferase gene in mammalian cells. J. Biolumin. Chemilumin. 11:31-37. Miyoshi, H., Blomer, U., Takahashi, M., Gage, F.H., and Verma, I.M. 1998. Development of a selfinactivating lentivirus vector. J. Virol. 72:81508157. Naldini, L., Blomer, U., Gallay, P., Ory, D., Mulligan, R., Gage, F.H., Verma, I.M., and Trono, D. 1996. In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263-267. Navarro-Galve, B., Villa, A., Bueno, C., Thompson, L., Johansen, J., and Martinez-Serrano, A. 2005. Gene marking of human neural stem/precursor cells using green fluorescent proteins. J. Gene Med. 7:18-29. Paroo, Z., Bollinger, R.A., Braasch, D.A., Richer, E., Corey, D.R., Antich, P.P., and Mason, R.P. 2004. Validating bioluminescence imaging as a high-throughput, quantitative modality for assessing tumor burden. Mol. Imaging 3:117124. Pichler, A., Prior, J.L., and Piwnica-Worms, D. 2004. Imaging reversal of multidrug resistance in living mice with bioluminescence: MDR1 P-glycoprotein transports coelenterazine. Proc. Natl. Acad. Sci. U.S.A. 101:1702-1707.

Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S., (eds.). 2009. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Rubio, F.J., Bueno, C., Villa, A., Navarro, B., and Martinez-Serrano, A. 2000. Genetically perpetuated human neural stem cells engraft and differentiate into the adult mammalian brain. Mol. Cell Neurosci. 16:1-13. Shah, K., Tung, C.H., Yang, K., Weissleder, R., and Breakefield, X.O. 2004. Inducible release of TRAIL fusion proteins from a proapoptotic form for tumor therapy. Cancer Res. 64:32363242. Shah, K., Bureau, E., Kim, D.E., Yang, K., Tang, Y., Weissleder, R., and Breakefield, X.O. 2005. Glioma therapy and real-time imaging of neural precursor cell migration and tumor regression. Ann. Neurol. 57:34-41. Shah, K., Hingtgen, S., Kasmieh, R., Figueiredo, J.L., Garcia-Garcia, E., Martinez-Serrano, A., Breakefield, X., and Weissleder, R. 2008. Bimodal viral vectors and in vivo imaging reveal the fate of human neural stem cells in experimental glioma model. J. Neurosci. 28:44064413. Snyder, E.Y. and Macklis, J.D. 1995. Multipotent neural progenitor or stem-like cells may be uniquely suited for therapy for some neurodegenerative conditions. Clin. Neurosci. 3:310316. Tang, Y., Shah, K., Messerli, S.M., Snyder, E., Breakefield, X., and Weissleder, R. 2003. In vivo tracking of neural progenitor cell migration to glioblastomas. Hum. Gene Ther. 14:1247-1254. Villa, A., Navarro-Galve, B., Bueno, C., Franco, S., Blasco, M.A., and Martinez-Serrano, A. 2004. Long-term molecular and cellular stability of human neural stem cell lines. Exp. Cell Res. 294:559-570.



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Functional Analysis of Adult Stem Cells Using Cre-Mediated Lineage Tracing

UNIT 5A.2

Diana L. Carlone1 1

Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts

ABSTRACT Lineage-tracing has been used for decades to establish cell fate maps during development. Recently, with the advent of genetic lineage-tracing techniques (employing Cre-lox recombination), it has been possible to permanently mark progenitor/stem cell populations within somatic tissues. In addition, pulse-chase studies have shown that only stem cells are capable of producing labeled progeny after an extensive period of chase. This unit focuses on the protocols used to target putative adult stem cells in vivo. Using these techniques, one should be able to functionally confirm or deny the stem cell capacity of C 2009 by John a given cell population. Curr. Protoc. Stem Cell Biol. 9:5A.2.1-5A.2.15. Wiley & Sons, Inc. Keywords: tamoxifen-inducible Cre recombination r lineage contribution r reporter activity r whole-mount analysis

INTRODUCTION Adult stem cells are elusive in many tissues. The promise of cell-based therapeutics for the treatment of human disease must first begin with the identification of functionally important stem cell populations. It is generally accepted that stem cells have the capacity for self-renewal and multi-lineage contribution within a given tissue. The Cre-lox system may be used to permanently mark cells of interest so that they can be observed for what they give rise to. If they give rise to no other lineages, they are not stem cells. If they give rise to multiple lineages, and subsequently are shown to self-renew, they are identified as stem cells. This unit focuses on lineage-tracing analysis using tamoxifen-inducible Cre-lox technology (Fig. 5A.2.1) to define the contribution of specific cell populations. This technique has been successfully employed to mark progenitor/stem cell populations in adult tissues. A detailed description of the Cre-lox system can be found elsewhere (Rossant and McMahon, 1999; Nagy, 2000; Branda and Dymecki, 2004). Briefly, Cre recombinase causes recombination of 34-bp loxP sequences and thus deletion of the intervening sequence (Fig. 5A.2.1). It is important to note that the orientation of the loxP sequences with respect to one another determines the recombination outcome (for a review, see Branda and Dymecki, 2004). Both loxP sequences must be in the same orientation for proper excision. Alteration in the orientation results in inversion of the intervening sequence and lack of deletion. While this technology is primarily used to induce tissue-specific knockout of genes in mice, when used to activate reporter genes it can indelibly mark discrete cell populations. The addition of inducible components such as the tamoxifeninducible Cre recombinase (CreER) to the system further allows for the study of temporal relationships between cell populations. Lineage tracing studies typically involve the use of double transgenic mice containing both a Cre-expressing transgene and a Cre reporter transgene. Cells are permanently marked during an initial Pulse and the contribution of their progeny to specific cell lineages is then determined during a period of Chase. Multiple strategies have been Current Protocols in Stem Cell Biology 5A.2.1-5A.2.15 Published online May 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a02s9 C 2009 John Wiley & Sons, Inc. Copyright


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A IoxP

ATAACTTCGTATAGCATACATTATACGAAGTTAT

B IoxP

gene X

IoxP

Cre recombinase

IoxP

Figure 5A.2.1 Schematic of the Cre-lox system. (A) The sequence for the loxP site is shown. The underlined sequence is the 8-bp core sequence where recombination occurs, and two flanking 13-bp inverted repeats. (B) Schematic of a transgene in which gene X is flanked by two loxP sites. In the presence of Cre recombinase the gene X is deleted leaving only a single loxP.

cell-specific promoter

CreERT X

stop LacZ

OR

Rosa26R reporter

LacZ

hPAP

Z/AP reporter

tamoxifen

LacZ

Cre-Mediated Lineage Tracing

hPAP

Figure 5A.2.2 Schematic of bigenic mouse model systems used for lineage tracing. To perform tracing studies, tamoxifen-inducible CreERT transgenic mice are crossed with either Rosa26R (left) or Z/AP (right) reporter mice. In the absence of ligand, β-galactosidase (LacZ) reporter is not expressed in the Rosa26R mice while Z/AP mice express LacZ. Upon administration of tamoxifen, recombination occurs in a cell-specific manner resulting in either LacZ (left) or human placental alkaline phosphatase (hPAP; right) expression. Once labeled, cells can then be chased to determine their contribution to distinct lineages.

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developed to indelibly mark cells for cell fate mapping. One strategy employs the use of a loxP-flanked dominant transcriptional stop sequence located upstream of a reporter gene. Following Cre-mediated deletion of this stop sequence, constitutive and permanent expression of the reporter gene is induced (see Fig. 5A.2.2; Rosa26R reporter mouse; Soriano, 1999). Another strategy involves the use of tandem reporter genes with the first gene flanked by loxP sites. Under baseline conditions, only the first gene is expressed constitutively. In this scheme, Cre-recombination removes the first gene, allowing the permanent expression of the second (see Fig. 5A.2.2; Z/AP reporter mouse; Lobe et al., 1999). In this unit, the protocols used for analysis of two commonly employed transgenes—β-galactosidase and alkaline phosphatase—are described.

WHOLE-MOUNT ANALYSIS OF β-GALACTOSIDASE ACTIVITY This protocol focuses on whole-mount analysis followed by immunohistochemistry to demonstrate that a discrete population of cells contributes to distinct differentiated lineages. To perform these studies, double transgenic mice containing both a tamoxifeninducible cell-specific CreER transgene and a Cre reporter transgene (β-galactosidase or alkaline phosphatase reporter) are used. As outlined in Critical Parameters, the choice of the Cre transgene as well as the reporter mouse line is dependent upon the scientific question being asked. An example of whole-mount analysis using the β-galactosidase reporter mouse line (Rosa26R) is illustrated in Figure 5A.2.3. While whole-mount analysis has the advantage of allowing for the detection of reporter activity in the context of the intact tissue, it does require subsequent histological analysis to confirm the identity of the marked cells. In this protocol, immunohistochemical analysis is performed using differentiation-specific antibodies to demonstrate the contribution of reporter-positive cells to specific cell lineages. Alternatively, if applicable, histological analysis using specific stains such as periodic acid-Schiff (PAS), which recognizes carbohydrates in tissue sections, can be used to demonstrate histologically that reporter-positive cells are differentiated (see Fig. 5A.2.3C-E).

BASIC PROTOCOL

NOTE: While validation of lineage contribution through immunohistochemical analysis of lacZ-stained regions is described, it is possible to co-label cells fluorescently using both differentiation-specific antibodies and β-galactosidase-specific antibodies using either paraffin or frozen sections.

Materials Tamoxifen-inducible Cre :: Rosa26R or Z/AP bigenic mice Tamoxifen or 4-hydroxytamoxifen (see recipe) Negative control mice (oil-treated bigenic or treated monogenic mice) Positive control mice (Rosa26; Jackson Laboratories cat. no. 002292) Phosphate-buffered saline, Ca++ - and Mg++ -free (CMF-PBS) LacZ fixative, wash, and staining buffers (see recipes) 32% (w/v) paraformaldehyde solution (EMS cat. no. 15714-S) 35% 70%, 80%, 90%, 95%, and 100% ethanol Xylene Paraffin 10 mM sodium citrate, pH 6.0 3% H2 O2 , optional Avidin and biotin blocking solutions (Vector Laboratories cat. no. SP-2001) Normal serum (species selection should match that of the secondary antibody; Sigma) Differentiation-specific antibodies Vectastain ABC Elite kit (species-specific kits are available dependent upon the primary antibody; Vector Laboratories)



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A

B

C

D

E

Figure 5A.2.3 Whole-mount and sectional analysis demonstrate lineage tracing in the small intestine. (A,B) Wholemount analysis of LacZ staining in the intestine following tamoxifen treatment (2-week chase) at low (A; 2×) and high (B; 11.25×) magnification. (C) Frozen sectional analysis of β-galactosidase activity in the small intestine following a 4-week chase. Tissue was subsequently stained with periodic acid-Schiff to detect lineage contribution. Magnification is 40×. (D,E) Histological analysis of lineage contribution by LacZ-positive cells in the small intestine following tamoxifen treatment. Following whole-mount analysis, LacZ-positive regions were paraffin embedded and sections were counterstained with periodic acid-Schiff. Co-labeling of LacZ and periodic acid-Schiff corresponds to goblet (D) and Paneth (E) cells. Magnification is 60×.

DAB substrate kit (Vector Laboratories cat. no. SK-4100) Nuclear Fast Red (Sigma cat. no. N3020) Cytoseal XYL mounting medium (Richard-Allan Scientific cat. no. 8312-4) 1.5-ml microcentrifuge tubes or 6-well tissue culture plates Platform shaker 37◦ C incubator Microtome Microscope slides Coplin jars Pressure cooker, microwave, or water bath Coverslips NOTE: Unless indicated, all steps in this protocol are performed at room temperature.

Induce Cre expression 1. Treat tamoxifen-inducible Cre :: Rosa26R bigenic mice with tamoxifen and control mice with oil. Collect tissue of interest after treatment (pulse) followed by a period of chase. Cre-Mediated Lineage Tracing

Bigenic mice are obtained by crossing tamoxifen-inducible Cre mice and commercially available β-galactosidase reporter Rosa26R mice (Jackson Laboratories cat. no. 003310).

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The dosage, route, and frequency of tamoxifen administration are dependent upon multiple factors, which are outlined in the Critical Parameters section and will need to be empirically determined for each Cre-expressing transgene and tissue of interest. In addition, the appropriate treatment should label single cells that can then be chased to determine their contribution to specific lineages. The length of chase is dependent upon multiple factors as described in Critical Parameters.

2. Wash tissue with ice-cold CMF-PBS to remove any contaminants and place into either a 1.5-ml microcentrifuge tube or 6-well tissue culture plate depending upon the size of tissue to be analyzed. Tissue from oil-treated bigenic or tamoxifen-treated monogenic mice should be used as a negative control to confirm specificity of the reaction. In addition, tissue from the Rosa26 mouse, which constitutively expresses β-galactosidase (Zambrowicz et al., 1997), can be used as a positive control. If necessary, tissues can be cleaned and washed for 5 to 10 min in cold CMF-PBS buffer containing 0.02% (v/v) NP-40 and 0.5 mM DTT prior to fixation. It has been found that this reduces background LacZ staining especially in whole-mount analysis of gastrointestinal tissues. The size/thickness of the tissue can affect the penetration of the staining solution thereby altering the efficiency of labeling. Therefore, using a small tissue biopsy, bisecting the tissue or, if necessary, gently poking holes into the tissue will increase the penetration. If, however, the entire tissue needs to be assayed, then analysis can be performed on tissue sections (see Alternate Protocol 2). Alternatively, if the tissue is thin enough, it can be processed intact while mounted on paraffin in tissue culture dishes using insect pins (Fine Science Tools). This approach is routinely used for whole-mount analysis of small intestine.

Fix tissues 3. Fix tissue in LacZ fixative solution for 1 hr on ice with shaking. To increase penetration and decrease background, the detergent NP-40 (0.02%) can be added to the fixative. Generally, a mild fixative such as glutaraldehyde is used; however, other fixatives can be used, including 4% (w/v) paraformaldehyde, or a combination of fixatives such as 0.2% (v/v) glutaraldehyde/2.0% (v/v) formaldehyde in CMF-PBS. CAUTION: Some fixatives can inhibit or diminish β-galactosidase activity; therefore, it may be necessary to test alternative fixatives.

4. Wash tissue three times with CMF-PBS for 10 min each on ice with shaking.

Stain for LacZ activity 5. Wash with LacZ wash buffer for 10 min on ice with shaking. 6. Incubate tissue in LacZ staining buffer from 1 to 24 hr at 37◦ C in the dark, until a dark color from the substrate reaction is seen, while the background is relatively unstained. The reaction is light sensitive so incubate sample in the dark. Generally, the reaction is stopped within 24 hr. The reaction can also be performed at lower temperatures such as 30o C, which has been shown to reduce background staining. However, a longer incubation time may be necessary.

7. To stop reaction, wash two to three times with CMF-PBS for 20 min each at room temperature with shaking. 8. To preserve tissue and staining, re-fix tissue with 4% paraformaldehyde in CMF-PBS for 1 to 2 hr (longer if necessary) at 4◦ C with shaking. 9. Place tissue in CMF-PBS and store at 4◦ C. Fixed tissue can be stored 3 to 6 months at 4◦ C without diffusion of the LacZ stain.



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Prepare tissues for immunohistochemical confirmation of lineages 10. Dehydrate whole-mount stained tissue through an ethanol series (35%, 70%, 90%, 100%) and xylene. Incubate the tissue two times in each solution, 1 hr each time. Embed in paraffin. The amount of time to dehydrate tissue may vary with the size of the tissue. Paraffin sections are used to confirm the contribution of a marked cell to distinct lineages because they allow better tissue histology than frozen sections.

11. Cut 4-μm sections and mount onto microscope slides. 12. Rehydrate sections through two changes of xylene, 3 min each, followed by two changes of 100% ethanol, 2 min each, and then through an ethanol series (95%, 90%, 80%, and 70%), 1 min each, using Coplin jars. All steps involving histological slides are performed using Coplin jars.

13. Wash with CMF-PBS for 5 min with shaking. For histological analysis of reporter positive cells, paraffin sections after rehydration can be counterstained with Nuclear Fast Red (see below).

Retrieve antigens 14. Perform antigen retrieval by boiling slides in 10 mM sodium citrate, pH 6.0, for 10 min using a pressure cooker, microwave, or water bath. Fixatives such as paraformaldehyde form protein cross-links that may mask antigenic sites giving negative (or weak) immunohistochemical results. Therefore, the antigen retrieval step unmasks the antigens/epitopes in paraffin sections. This buffer is commonly used and works well with most antibodies.

15. Allow slides to cool in buffer for 45 to 60 min. 16. Wash slides two to three with CMF-PBS, 5 min each, with shaking.

Treat to reduce background 17. (Optional) Incubate slides in 3% H2 O2 for 15 min with shaking. This step blocks endogenous peroxidase activity. Because not all tissues exhibit endogenous activity, this step is optional depending upon the tissue of interest.

18. Wash slides three times with CMF-PBS, 5 min each, with shaking. 19. Incubate slides with avidin and biotin blocking solutions, 15 min each, with shaking. Wash three times with CMF-PBS in between each step. Like peroxidase activity, some tissues exhibit endogenous biotin activity; therefore, this step is also optional depending upon the tissue of interest.

20. To reduce non-specific background, block sections with 1% to 5% normal serum in CMF-PBS for 15 to 30 min with shaking. The exact percentage of serum to be used needs to be determined empirically for each antibody. In addition, the normal serum used should be from the same species as the secondary antibody. If necessary, 0.1% to 0.3% Triton X-100 can be used in the blocking solution to further decrease background.

Immunostain slides 21. Incubate sections with primary antibody at appropriate dilution in blocking solution for 1 hr at room temperature to overnight at 4◦ C. The exact incubation conditions for each antibody must be empirically determined. Cre-Mediated Lineage Tracing

22. Wash slides three times with CMF-PBS, 5 min each, with shaking.

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23. Incubate with biotinylated secondary antibody (Vectastain Elite ABC kit) at appropriate dilution in blocking solution per the manufacturer’s instructions for 30 min at 37◦ C. Selection of appropriate Vectastain Elite ABC kit is dependent upon the species used to generate the primary antibody.

24. Wash slides three times with CMF-PBS, 5 min each, with shaking.

Detect antibody binding 25. Incubate sections with ABC reagent 30 min at 37◦ C. 26. Wash slides three times with CMF-PBS, 5 min each, with shaking. 27. Incubate slides in 3,3 -diaminobenzidine (DAB) substrate solution for 1 to 5 min. The DAB solution yields a brown substrate color. Alternatively, the VIP substrate solution (Vector Laboratories cat. no. SK-4600) can be used and yields a purple substrate color. In addition, the precise incubation time must be determined empirically for each antibody.

28. To stop reaction, wash slides with water for 5 min with shaking. 29. If necessary, counterstain sections with Nuclear Fast Red solution for 30 sec to 1 min. This solution is light sensitive. Nuclear Fast Red is diluted 1:1 with distilled water, filtered, and stored for 3 to 6 months at room temperature. This stain can be reused multiple times and re-filtered as needed.

30. Dehydrate slides for 2 min each using 70%, 90%, and 100% ethanol and xylene. 31. Mount slides with coverslips using Cytoseal XYL mounting medium.

WHOLE-MOUNT ANALYSIS OF ALKALINE PHOSPHATASE ACTIVITY Although many researchers use LacZ staining to permanently trace the contribution of stem cells, Cre/lox reporter mice utilizing other reporters such as the enzyme, alkaline phosphatase, can also be used in whole-mount analysis. This protocol can therefore be used as an alternative approach to define the role of a discrete cell population as stem cells using lineage tracing technology.


Additional Materials (also see Basic Protocol) Alkaline phosphatase fixative solution (see recipe) AP buffer (see recipe) BM Purple AP substrate (Roche Diagnostics cat. no. 11 442 074 001) PTM buffer (see recipe) 70◦ to 75◦ C incubator Collect tissue 1. Treat tamoxifen-inducible Cre :: Z/AP bigenic mice with tamoxifen and collect tissue at pulse and chase time points. As indicated in the Basic Protocol, tissue from oil-treated bigenic or tamoxifen-treated monogenic mice should be used as a negative control. In addition, constitutively expressing alkaline phosphatase mice can be used as a positive control. If appropriate, alkaline phosphatase and LacZ can be detected in the same tissue. Staining for β-galactosidase must be performed first due to its sensitivity to heat, which is used to reduce endogenous alkaline phosphatase activity (see step 4). After X-gal staining, tissue should be rinsed well with CMF-PBS prior to performing the alkaline phosphatase protocol.



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Fix tissue 2. Fix tissue with alkaline phosphatase fixative solution for 30 min on ice with shaking. 3. Wash tissue three times with CMF-PBS, 10 min each, on ice with shaking. 4. Heat-inactivate endogenous alkaline phosphatase activity by incubating in CMF-PBS 30 min at 70◦ to 75◦ C. 5. Wash with CMF-PBS 10 min at room temperature with shaking.

Detect AP activity 6. Wash with AP buffer 10 min at room temperature with shaking. 7. Stain with BM Purple AP substrate up to 36 hr at 4◦ C, until a dark color from the substrate reaction is seen. Incubation at room temperature will accelerate the reaction but may result in diffusion of the stain.

8. Wash tissue three times with PTM buffer, 10 min each, at room temperature with shaking. 9. To preserve the staining, re-fix the tissue with 4% paraformaldehyde in CMF-PBS for 1 to 2 hr (longer if necessary) at 4◦ C with shaking. 10. Place tissue in CMF-PBS and store for 3 to 6 months at 4◦ C. For immunohistochemical analysis to confirm lineage contribution, tissues can be processed similar to LacZ-stained whole-mount tissue, see Basic Protocol, step 10. ALTERNATE PROTOCOL 2

SECTIONAL ANALYSIS FOR β-GALACTOSIDASE OR ALKALINE PHOSPHATASE ACTIVITY Although whole-mount analysis allows for reporter detection in the intact tissue, analysis of tissue sections for reporter activity identifies the specific marked cell histologically. This protocol can therefore be used as an alternative approach or, in many instances, in combination with whole-mount analysis to further define the role of a discrete cell population as stem cells using lineage tracing technology (see Fig. 5A.2.3).

Additional Materials (also see Basic Protocol) Tissue-Tek OCT (Sakura, cat. no. 4583) 0.2% glutaraldehyde/2 mM MgCl2 in CMF-PBS 0.2% glutaraldehyde in CMF-PBS AP buffer (see recipe) 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium (BCIP/NBT) solution (see recipe or Vector Laboratories cat. no. SK-5400) Cryomolds Cryostat 70◦ to 75◦ C incubator Analyze sections for reporter activity 1. Isolate tissue following chase and pulse time points as determined in whole-mount analysis and place directly into cryomolds containing Tissue-Tek OCT and freeze on dry ice. Alternatively, tissues can be fixed in 4% paraformaldehyde for 1 to 2 hr on ice, incubated in 0.6 M sucrose overnight at 4o C, and then embedded in OCT. Sucrose acts as a cryoprotectant to minimize ice crystal damage, allowing for better microscopic morphology. Cre-Mediated Lineage Tracing

The heating process used during the embedding of tissue for paraffin sections inactivates β-galactosidase; therefore, all sectional analysis is performed using frozen sections.

5A.2.8 Supplement 9


2. Section tissue (10-μm), dry up to 2 hr at room temperature, and store at −20◦ C. Stain for β-galactosidase activity 3a. Fix slides in 0.2% glutaraldehyde/2 mM MgCl2 in CMF-PBS for 10 min on ice with shaking. Alternative fixatives such as 4% paraformaldehyde may be used; however, as indicated with the whole-mount analysis, reporter activity may vary with harsher fixatives.

4a. Wash slides two to three times with LacZ wash buffer, 10 min each, at room temperature with shaking. 5a. Incubate in LacZ staining buffer 1 to 24 hr at 37◦ C in the dark. 6a. To stop the reaction, wash slides three times with CMF-PBS, 10 min each, followed by a quick rinse in water at room temperature. 7a. Counterstain sections with Nuclear Fast Red solution for 30 sec to 1 min. 8a. Dehydrate slides for 2 min each using 70%, 90%, and 100% ethanol and xylene. 9a. Mount slides with coverslips using Cytoseal XYL mounting medium. Stain sections for alkaline phosphatase activity 3b. Fix slides in 0.2% glutaraldehyde in CMF-PBS for 10 min on ice with shaking. 4b. Wash slides three times with CMF-PBS, 5 min each, at room temperature with shaking. 5b. Inactivate endogenous alkaline phosphatase by incubating slides in CMF-PBS for 30 min at 70◦ to 75◦ C. 6b. Wash slides with CMF-PBS for 10 min at room temperature with shaking. 7b. Wash slides with AP buffer for 10 min at room temperature with shaking. 8b. Stain slides with BCIP/NBT solution for 10 to 30 min at room temperature. Staining solution should be placed directly onto the slides. Sensitivity of substrate can be increased by lengthening the incubation time.

9b. Wash slides in CMF-PBS and process as indicated above in steps 7a through 9a.

REAGENTS AND SOLUTIONS For all solutions, use deionized, distilled water or equivalent in recipes and protocol steps. Unless indicated, all solutions are made up in water. Suppliers for non-common chemicals are indicated.

Alkaline phosphatase fixative solution 0.2% (v/v) glutaraldehyde 50 mM EGTA, pH 7.3 100 mM MgCl2 0.02% (v/v) NP-40 0.01% (w/v) sodium deoxycholate Prepare fresh To increase the penetration of alkaline phosphatase (AP) substrates, 0.02% NP-40 and 0.01% sodium deoxycholate have been added to the fixative.

AP buffer 10 mM Tris·Cl, pH 9.5 100 mM NaCl continued



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10 mM MgCl2 Store up to 1 year at room temperature LacZ fixative solution 0.2% (v/v) glutaraldehyde 2 mM MgCl2 CMF-PBS Prepare fresh LacZ staining buffer 0.5 mg/ml X-gal (stock is 40 mg/ml in dimethylformamide) 5 mM K3 Fe(CN)6 5 mM K4 Fe (CN)6 -3H2 O LacZ wash buffer (see recipe) Prepare fresh Staining buffer should be made fresh each time. Use CMF-PBS at approximately pH 7.4 for all steps. If necessary, use CMF-PBS at pH 8.0 for LacZ staining buffer to decrease background β-galactosidase activity.

LacZ wash buffer 2 mM MgCl2 0.01% (w/v) deoxycholate 0.02% (v/v) NP-40 CMF-PBS Store up to 1 year at room temperature NBT/BCIP stain solution 100 mM Tris·Cl, pH 9.5 100 mM NaCl 50 mM MgCl2 0.01% (w/v) sodium deoxycholate 0.02% (v/v) NP-40 337 μg/ml nitroblue tetrazolium salt (NBT; Sigma cat. no. N6876) 175 μg/ml 5-bromo-4-chloro-3-indolyl phosphate, disodium salt (BCIP; Sigma cat. no. B1026) Prepare fresh PTM buffer 0.1% (v/v) Tween-20 2 mM MgCl2 CMF-PBS Store up to 1 year at room temperature Tamoxifen or 4-hydroxytamoxifen Resuspend tamoxifen (Sigma cat. no. T5648) or 4-hydroxytamoxifen (70% Z isomer, 30% E isomer 4-OHT; Sigma cat. no. H6278) at a concentration of 10 to 20 mg/ml in peanut oil (Indra et al., 1999; Kühbandner et al., 2000). Add 500 μl of 100% ethanol to 100 mg of tamoxifen followed by 9.5 ml of peanut oil. Dispense into aliquots and store for up to 4 weeks at −20◦ C. Corn oil can also be used to resuspend tamoxifen or 4-OHT. Cre-Mediated Lineage Tracing

Tamoxifen is fairly soluble while 4-OHT requires sonication or heating. Both will precipitate at cold temperatures; therefore, it may be necessary to heat or resonicate prior to use.



COMMENTARY Background Information The identification of tissue progenitor/stem cells requires demonstrating both their selfrenewal potential and their capacity to contribute to multiple cell lineages within a tissue. While current studies use genetic models to experimentally demonstrate stem cell function, the scientific literature is full of alternate and historical approaches to map the fate of a cell (reviewed in Stern and Fraser, 2001). Early studies focused on deciphering cell lineage using a variety of techniques to follow cells and their descendants. For example, endogenous expression of alkaline phosphatase by germ cells allowed for tracing of these cells throughout development (Chiquoine, 1954). In addition, direct visualization by the use of pigmentation differences among cells has allowed for the construction of lineage trees in developing organisms, including the complete cell fate map of the nematode, Caenorhabditis elegans (Sulston et al., 1983; Thomas et al., 1996). This technique is limited however by an inability to identify and trace single cells over time and the problems inherent to increasingly opaque embryos. To combat these problems, a variety of approaches have been taken to mark cells. The use of vital dyes to trace living cells was attempted but was found to be ineffective due to their water solubility, which resulted in transfer of dye to unrelated cells. Eventually, multicolored carbocyanine dyes were generated, which are lipid soluble/water insoluble and localize within the cell membrane (Axelrod, 1979; Serbedzija et al., 1989; YablonkaReuveni, 1989; Eagleson and Harris, 1990). While cells readily take up these dyes, it proved difficult to mark single cells, thus this technique has been used to track the fate of cell populations. Additional strategies have included the use of radiolabeled compounds to mark cells prior to introduction into embryos as well as the generation of interspecies chimeras (e.g., chick/quail), which relies upon species-specific differences in cell pigmentation and size to distinguish between donor and host cells (Le Douarin, 1973; Dupin et al., 1998). Marking and tracing of single cells in both vertebrates and invertebrates have been accomplished by single-cell injection with inert tracers such as the enzyme horseradish peroxidase, or fluorescently-labeled compounds such as dextran or lysine (Weisblat et al., 1978; Lawson et al., 1986; Gimlich and Braun, 1985;

Peralta and Denaro, 2003). While technically challenging, very elegant cell fate mapping is possible with such approaches. The biggest disadvantage is that the tracer becomes diluted in dividing cells. To determine the fate of cells that might migrate during development, colloidal goldlabeled monoclonal antibodies were developed and used to track the descendant of cells that express a common surface antigen regardless of their position in the embryo (Stern and Canning, 1990). Antigen-expressing cells internalize the antibody/gold complex and pass it on to their descendants. However, the marker can only be detected for a few divisions. To overcome this problem, several groups developed retroviral vectors that would label cells through the introduction of marker genes, e.g., alkaline phosphatase or β-galactosidase (Cepko et al., 1984; Sanes et al., 1986). Diluted viral stock solutions, as well as replicationdeficient strains, were subsequently employed to increase the probability that marked cells would be clonally derived. This approach has been used to trace cell lineages in the nervous system (Cepko et al., 1984; Price et al., 1987). Given that viral targeting is not celltype-specific, these lineage-mapping studies must be performed retrospectively. In addition, it is impossible to rule out that adjacent cells were not also initially labeled via an independent infection event. To address this, complex retroviral libraries were generated (Golden et al., 1995) with the belief that the infection was a random event and thus it would be highly unlikely that adjacent cells would be infected by the same retrovirus. Distinction between the various retroviruses could be confirmed by PCR. Although this approach increased the rigor, the analysis remained retrospective and required that each cell type be analyzed by PCR to determine their relatedness to neighboring labeled cells. Therefore, this method required the ability to isolate the cell types of interest. Alternatively, spontaneous DNA recombination events (+/− mutagens) have been used to retrospectively trace cells via activation of marker expression in transgenic mice (Bonnerot and Nicolas, 1993; Bjerknes and Cheng, 1999). Once labeled, cells are transplanted into adult animals to demonstrate cell fate mapping. The contribution of a putative stem cell to various cell lineages is assessed through analysis of marker expression or, alternatively, identification of the Y chromosome when



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male donor cells were injected into female recipients. This approach has been used to study hematopoietic stem engraftment and multi-lineage contribution. While these approaches have yielded important insight into cell fate, increasingly, the use of site-specific recombinase systems such as Cre-lox has become the gold standard for cell lineage studies. This technology allows for the control over the temporal and spatial marking of selective cell populations through the use of gene-specific promoters to regulate recombinase expression (for review see Sauer, 1998; Nagy, 2000; Branda and Dymecki, 2004). To regulate the onset of Cre expression independent from endogenous regulatory elements, inducible Cre recombinases have been generated that allow for refined labeling of specific single cells in response to ligand administration (Furth et al., 1994; Feil et al., 1996; Kellendonk et al., 1996; Rivera et al., 1996; Brocard et al., 1998; Danielian et al., 1998; Utomo et al., 1999; Schönig et al., 2002). Specifically, in the tamoxifen-inducible Cre-lox system (used in this unit), Cre recombinase (CreER) is fused to a mutated ligand-binding domain of the estrogen receptor, which specifically binds tamoxifen and not the endogenous estrogen. In the absence of ligand, CreER is retained within the cytoplasm. Upon tamoxifen administration, the CreER protein translocates to the nucleus where it excises the loxP-targeted site. Furthermore, the commercial availability of a variety of Cre-reporter mouse lines, which upon recombination express colorimetric, enzymatic, or fluorescent proteins, allows for greater diversity in the use of lineage-tracing to define stem cells.



While the lineage-tracing approach can demonstrate “stemness” of a particular cell population, it does require some a priori knowledge. First, one must determine whether a specific gene selectively marks the cell of interest. Second, determine whether a Cre mouse line already exists in which the promoter of the gene of interest regulates Cre recombinase expression. Preferably this line should be inducible, allowing for controlled temporal marking of the cell. Third, determine the Cre reporter mouse line to be used. The choice of which Cre reporter line to use is dependent upon several things including whether the reporter is expressed in the cell and tissue of interest as well as the type of analysis that

will be performed. Although most reporters are considered to be ubiquitously expressed, it has been found that is not always the case; therefore, when possible, confirming the expression of the reporter in the cell of interest is optimal. In addition, while this unit focuses on β-galactosidase and alkaline phosphatase reporters, fluorescent Cre reporter mouse lines have been used to trace lineage contribution. Generally, colorimetric and enzymatic reporters are convenient to use for confirming lineage contribution by a stem cell; however, fluorescent reporters have the added advantage of allowing for single-cell isolation and analysis via flow cytometry. The following Websites contain additional information on the various Cre and Cre reporter mouse lines available for lineage tracing: The Jackson Laboratory: http://www.jax.org or Dr. Andras Nagy’s laboratory Website: http://www.mshri.on.ca/nagy. Although stem cells have been identified in tissues that are highly regenerative such as the skin, intestine, and blood, adult stem cells in other tissues may require some type of regenerative stimulus such as injury to awaken them. Therefore, prior knowledge of the mechanism(s) involved in stem cell activation within the tissue of interest will be advantageous before performing lineage tracing studies. Administration of the inducing agent varies depending upon the model system used. In the case of the tamoxifen-inducible Cre mouse models, tamoxifen can be administration intraperitoneally (i.p.), subcutaneously (s.c.), and orally (p.o.) as well as by pellet implants. The amount and length of administration is highly dependent upon the strength and expression pattern of the promoter regulating the Cre recombinase and will have to be determined empirically by each researcher. Tamoxifen is generally given at a dose of 1 to 2 mg/day i.p. for up to 5 days. Analysis of reporter activity is determined at the end of the pulse period (1 to 3 days after the last injection) at which time cell-specific labeling should be detected. It is important to note that this regimen should be used as a starting point and modifications in either length of delivery and/or dosage may be necessary depending upon the tissue of interest. If weak or no labeling is detected, then increasing the dose of tamoxifen may be warranted. Alternatively, 4-hydroxytamoxifen (4-OHT), a metabolite of tamoxifen, which exhibits a more potent activity, may be used instead to increase labeling. However, tamoxifen is often preferred over 4OHT because it is more soluble in solution and less costly. It is worth noting that high



levels of tamoxifen or 4-OHT administered i.p. can have deleterious affects on mice. Varying the timing or mode of administration can circumvent this problem, e.g., administering i.p. injections every other day instead of daily or injecting s.c. or p.o. instead of i.p. Once labeled, the time period in which to analyze the cell’s contribution to various progeny (chase) must be determined. The exact time is dependent upon the tissue of interest as well as whether the stem cell population requires a regenerative stimulus to induce new lineage development. For a general reference, the chase can range from weeks to months. As indicated in the Basic Protocol, contribution of the marked stem cell to differentiated progeny can be confirmed by co-localization of the reporter with differentiation-specific markers by immunohistochemistry or immunofluorescence (depending upon the reporter). This approach however relies upon the availability of specific antibodies. If none are available, alternative approaches may be used such as differentiation-specific gene marker analysis of isolated reporter-positive cells using flow cytometry. As indicated earlier, multiple fluorescent reporter mouse lines are available as well as commercially available flow cytometric kits for β-galactosidase activity (Invitrogen). High background LacZ staining may be due to endogenous enzymatic activity, which has been reported in a variety of tissues. Endogenous β-galactosidase is normally active at low pH (∼4) while bacterial β-galactosidase is active at a more neutral pH. Therefore, increasing the pH of CMF-PBS in the LacZ wash and staining buffers should decrease background LacZ staining. CMF-PBS at a pH between 7.4 and 8.0 is usually used. Extreme alkaline conditions (pH 8.5 to 9.0), however, can inhibit bacterial β-galactosidase activity. In addition, the incubation temperature and time can affect background staining. Minimizing the reaction time to 2 hr and reducing the temperature from 37◦ C to room temperature dramatically reduces background staining. It is important to note that the optimal staining conditions for each tissue needs to be determined empirically using both positive and negative control tissues. There are several possible explanations for a lack of or low reporter activity. As a general rule, tissues from positive control mice are used as a control to validate that the assay is functioning correctly. Absence of reporter activity may be due to either the re-

porter or the Cre recombinase not being expressed in the cell type of interest. To confirm the cellular specificity of Cre recombinase expression, in situ hybridization or immunohistochemical analysis may be performed. As indicated above, although most reporters are believed to be globally expressed upon recombination, variegated expression both across tissues and within select tissues has been found. Therefore, confirmation of reporter expression in the cell and tissue of interest is essential before performing lineage tracing experiments. This can be done by assessing reporter activity in a positive control mouse such as Rosa26 (Zambrowicz et al., 1997), which expresses β-galactosidase from the same genomic locus as the Cre reporter mouse line, suggesting comparable control elements. Alternatively, decreased penetration may affect the staining efficiency in whole-mount analysis. This can be addressed by either analyzing smaller tissue pieces, addition of detergents, or alternatively performing sectional analysis. Finally, low levels of reporter activity may be due to low efficiency of recombination. The strength of the promoter and the number of cells expressing the gene of interest can affect the efficiency of labeling. Increasing the dosage and/or altering the administration regimen may yield higher recombination efficiencies. Alternatively, it may be necessary to use a different inducible system such as the doxycycline system (Schönig et al., 2002) to increase the relative efficiency.

Anticipated Results In response to tamoxifen administration, reporter activity should be detected in a small population of cells preferably single cells at the pulse time point. If multiple cells are labeled, then decreasing the tamoxifen dosage or length of administration should yield single cells. Depending upon the tissue, wholemount analysis of single cells may be difficult; therefore, confirmation of single cells may require sectional analysis. Following this, if a cell functions as a stem cell, then contribution of label to other cell types including differentiated cells within a defined time period should be observed.

Time Considerations The time period required to detect lineage contribution depends upon a number of factors including the regenerative nature of the tissue. For example, the intestine is highly regenerative and turns over every 4 to 5 days; therefore,



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it would require a relatively short chase period to detect stem cells. Alternatively, a more quiescent stem cell population may take much longer to be activated and would therefore require an extended time period before lineage contribution would be detected. In addition, it is important to note that the requirement of inducing regeneration by tissue injury (thereby activating a stem cell population) can further extend the chase time period. Thus, a general time-frame for conducting these experiments relies upon the tissue of interest and will have to be determined by each researcher. It may take a chase weeks to months to detect a stem cell contribution.

Literature Cited Axelrod, D. 1979. Carbocyanine dye orientation in red cell membrane studied by microscopic fluorescence polarization. Biophys. J. 26:557573.

Gimlich, R.L. and Braun, J. 1985. Improved fluorescent compounds for tracing cell lineage. Dev. Biol. 109:509-514. Golden, J.A., Fields-Berry, S.C., and Cepko, C.L. 1995. Construction and characterization of a highly complex retroviral library for lineage analysis. Proc. Natl. Acad. Sci. U.S.A. 92:57045708. Indra, A.K., Warot, X., Brocard, J., Bornert, J.M., Xiao, J.H., Chambon, P., and Metzger, D. 1999. Temporally controlled site-specific mutagenesis in the basal layer of the epidermis: Comparison of the recombinase activity of the tamoxifeninducible Cre-EFT and Cre-ERT2 recombinases. Nucleic Acids Res. 27:4324-4327.

Bjerknes, M. and Cheng, H. 1999. Clonal analysis of mouse intestinal epithelial progenitors. Gastroenterology 116:7-14.

Kellendonk, C., Tronche, F., Monaghan, A.-P., Angrand, P.-O., Stewart, F., and Schutz, G. 1996. Regulation of Cre recombinase activity by the synthetic steroid RU486. Nucleic Acids Res. 24:1404-1411.

Bonnerot, C. and Nicolas, J.F. 1993. Clonal analysis in the intact mouse embryo by intragenic homologous recombination. C.R. Acad. Sci. III 316:1207-1217.

Kühbandner, S., Brummer, S., Metzger, D., Chambon, P., Hofmann, F., and Feil, R. 2000. Temporally controlled somatic mutagenesis in smooth muscle. Genesis 28:15-22.

Branda, C.S. and Dymecki, S.M. 2004. Talking about a revolution: The impact of site-specific recombinase on genetic analyses in mice. Dev. Cell 6:7-28.

Lawson, K.A., Meneses, J.J., and Pedersen, R.A. 1986. Cell fate and cell lineage in the endoderm of the presomite mouse embryo, studied with an intracellular tracer. Dev. Biol. 115:325-339.

Brocard, J., Feil, R., Chambon, P., and Metzger, D. 1998. A chimeric Cre recombinase inducible by synthetic, but not natural ligands of glucocorticoid receptor. Nucleic Acid Res. 26:40864090.

Le Douarin, N. 1973. A biological cell labeling technique and its use in experimental embryology. Dev. Biol. 30:217-222.

Cepko, C.L., Roberts, B.E., and Mulligan, R.C. 1984. Construction and applications of a highly transmissible murine retrovirus shuttle vector. Cell 37:1053-1062. Chiquoine, A.D. 1954. The identification, origin, and migration of the primordial germ cells in the mouse embryo. Anat. Rec. 118:135-146. Danielian, P.S., Muccino, D., Rowitch, D.H., Michael, S.K., and McMahon, A.P. 1998. Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible form of Cre recombinase. Curr. Biol. 8:1323-1326. Dupin, E., Ziller, C., and Le Douarin, N.M. 1998. The avian embryo as a model in developmental studies: Chimeras and in vitro clonal analysis. Curr. Top. Dev. Biol. 36:1-35. Eagleson, G.W. and Harris, W.A. 1990. Mapping of the presumptive brain regions in the neural plate of Xenopus laevis. J. Neurobiol. 21:427-440.


Furth, P.A., St. Onge, L., Böger, H., Gruss, P., Gossen, M., Kistner, A., Bujard, H., and Hennighausen, L. 1994. Temporal control of gene expression in transgenic mice by a tetracycline-responsive promoter. Proc. Natl. Acad. Sci. U.S.A. 91:9302-9306.

Feil, R., Brocard, J., Mascrez, B., LeMeur, M., Metzger, D., and Chambon, P. 1996. Ligand-activated site-specific recombination in mice. Proc. Natl. Acad. Sci. U.S.A. 93:1088710890.

Lobe, C.G., Koop, K.E., Kreppner, W., Lomeli, H., Gertsenstein, M., and Nagy, A. 1999. Z/AP, a double reporter for Cre-mediated recombination. Dev. Biol. 208:281-292. Nagy, A. 2000. Cre recombinase: The universal reagent for genome tailoring. Genesis 26:99109. Peralta, M. and Denaro, F.J. 2003. The horseradish peroxidase technique for cell lineage studies. Cell Mol. Biol. 49:1371-1375. Price, J., Turner, D., and Cepko, C. 1987. Lineage analysis in the vertebrate nervous system by retrovirus-mediated gene transfer. Proc. Natl. Acad. Sci. U.S.A. 84:156-160. Rivera, V.M., Clackson, T., Natesan, S., Pollock, R., Amara, J.F., Keenan, T., Magari, S.R., Phillips, T., Courage, N.L., Cerasoli, F., Hot, D.A., and Gilman, M. 1996. A humanized system for pharmacological control of gene-expression. Nat. Med. 2:1028-1032. Rossant, J. and McMahon, A. 1999. “Cre”-ating mouse mutants: A meeting review on conditional mouse genetics. Genes Dev. 13:142145. Sanes, J.R., Rubenstein, J.L.R., and Nicolas, J-F. 1986. Use of a recombinant retrovirus to



study post-implantation cell lineage in mouse embryos. EMBO J. 5:3133-3142. Sauer, B. 1998. Inducible gene targeting in mice using the cre/lox system. Methods 14:381-392.

Thomas, C., DeVries, P., Hardin, J., and White, J. 1996. Four-dimensional imaging: Computer visualization of 3D movements in living specimens. Science 273:603-607

Schönig, K., Schwenk, F., Rajewsky, K., and Bujard, H. 2002. Stringent doxycycline dependent control of CRE recombinase in vivo. Nucleic Acids Res. 30:e134.

Utomo, A.R., Nikitin, A.Y., and Lee, W.H. 1999. Temporal, spatial, and cell type-specific control of Cre-mediated DNA recombination in transgenic mice. Nat. Biotechnol. 17:1091-1096.

Serbedzija, G.N., Bronner-Fraser, M., and Fraser, S.E. 1989. A vital dye analysis of the timing and pathways of avian trunk neural crest cell migration. Development 106:809-816.

Weisblat, D.A., Sawyer, R.T., and Stent, G.S. 1978. Cell lineage analysis by intracellular injection of a tracer enzyme. Science 202:1295-1298.

Soriano, P. 1999. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat. Genet. 21:70-71. Stern, C.D. and Canning, D.R. 1990. Origin of cells giving rise to mesoderm and endoderm in chick embryo. Nature 343:273-275. Stern, C.D. and Fraser, S.E. 2001. Tracing the lineage of tracing cell lineages. Nat. Cell Biol. 3:E216-E218. Sulston, J.E., Schierenberg, E., White, J.G., and Thomson, J.N. 1983. The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev. Biol. 100:64-119.

Yablonka-Reuveni, Z. 1989. The emergence of the endothelial cell lineage in the chick embryo can be detected by uptake of acetylated low density lipoprotein and the presence of a von Willebrand-like factor. Dev. Biol. 132:230240. Zambrowicz, B.P., Imamoto, A., Fiering, S., Herzenberg, L.A., Ker, W.G., and Soriano, P. 1997. Disruption of overlapping transcripts in the ROSA βgeo 26 gene trap strain leads to widespread expression of β-galactosidase in mouse embryos and hematopoietic cells. Proc. Natl. Acad. Sci. U.S.A. 94:3789-3794.



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Magnetic Resonance Imaging of Human Embryonic Stem Cells

UNIT 5A.3

Jaehoon Chung,1 Mayumi Yamada,1 and Phillip C. Yang1 1

Stanford University School of Medicine, Stanford, California

ABSTRACT Magnetic resonance imaging (MRI) may emerge as an ideal non-invasive imaging modality to monitor stem cell therapy in the failing heart. This imaging modality generates any arbitrary tomographic view at high spatial and temporal resolution with exquisite intrinsic tissue contrast. This capability enables robust evaluation of both the cardiac anatomy and function. Traditionally, superparamagnetic iron oxide nanoparticle (SPIO) has been widely used for cellular MRI due to SPIO’s ability to enhance sensitivity of MRI by inducing remarkable hypointense, negative signal, “blooming effect” on T2*-weighted MRI acquisition. Recently, manganese chloride (MnCl2 ) has been reported by our laboratory for its ability as a contrast agent to track biological activity of viable cells. Hyperintense, positive signals can be achieved from the Mn2+ -labeled stem cells on T1-weighted MRI acquisition. Cytotoxicity is a potential drawback of Mn2+ labeling of the cells. However, in our laboratory the labeling method has been optimized to minimize cytotoxic effects. This article describes two different magnetic labeling methods of human embryonic stem cells (hESC) using SPIO and MnCl2 . Curr. C 2009 by John Wiley & Sons, Inc. Protoc. Stem Cell Biol. 10:5A.3.1-5A.3.9. Keywords: human embryonic stem cell (hESC) r magnetic resonance imaging (MRI) r superparamagnetic iron oxide (SPIO) r manganese

INTRODUCTION Magnetic resonance imaging (MRI) may emerge as an ideal non-invasive imaging modality to monitor stem cell therapy in the failing heart. This imaging modality generates any arbitrary tomographic view at high spatial and temporal resolution with exquisite intrinsic tissue contrast. This capability enables robust evaluation of both the cardiac anatomy and function. Traditionally, superparamagnetic iron oxide nanoparticle (SPIO) has been widely used for cellular MRI due to SPIO’s ability to enhance sensitivity of MRI by inducing remarkable hypointense, negative signal, “blooming effect” on T2*-weighted MRI acquisition (Fig. 5A.3.1). Recently, manganese chloride (MnCl2 ) has been reported by our laboratory for its ability as a contrast agent to track biological activity of viable cells. Hyperintense, positive signals could be achieved from the Mn2+ -labeled stem cells on T1-weighted MRI acquisition (Fig. 5A.3.2). Cytotoxicity was a potential drawback of Mn2+ labeling of the cells but in our laboratory, the labeling method has been optimized to minimize cytotoxic effects. This article describes two different magnetic labeling methods of human embryonic stem cells (hESC) using SPIO (Basic Protocol) and MnCl2 (Alternate Protocol). NOTE: All procedures must be performed in a sterile cell culture hood. All solutions and equipment in contact with live cells must be sterile, and aseptic technique must be used accordingly. NOTE: All culture incubations must be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Genetic Manipulation of Stem Cells Current Protocols in Stem Cell Biology 5A.3.1-5A.3.9 Published online August 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a03s10 C 2009 John Wiley & Sons, Inc. Copyright


a

* c b

e

d

Figure 5A.3.1 Magnetic resonance imaging of SPIO-labeled human embryonic stem cells at a coronal view. The different quantities of SPIO-labeled hESC (a) 2 million, (b) 1 million, (c) 0.5 million, (d) 0.1 million, (e) 0.05 million, and control (non-labeled designated with an *).

i

ii

iii

iv

v

vi

vii

Figure 5A.3.2 3 × 106 MnCl2 -labeled hESC at an axial view using the following concentrations: (i) control [0.9% (w/v) sodium chloride solution only], (ii) 0.01 mM, (iii) 0.05 mM, (iv) 0.10 mM, (v) 0.50 mM, (vi) 1.00 mM, and (vii) 3.00 mM. Dose-appropriate increase in T1-weighted positive contrast is seen up to 1.00 mM. hESC are indicated by a black arrow.

BASIC PROTOCOL

DIRECT MAGNETIC LABELING OF HUMAN EMBRYONIC STEM CELLS (hESC) USING SPIO Direct labeling of hESC with SPIO is a convenient and robust method. This technique requires the addition of transfection agents such as protamine sulfate, poly-L-lysine (PLL) or electroporation to increase efficiency of labeling (Frank et al., 2002; Suzuki et al., 2007). Here we introduce a protocol using protamine sulfate (PS). The dextrancoated SPIO particles carry a negative charge. By coating SPIO with positively charged PS, neutral or slightly positive electrical charge can be formed on the surface of the SPIO-PS complex. Consequently, electrostatic interaction between the cell membrane and SPIO-PS complex enhances cellular uptake of SPIO-PS complex via mechanisms including endocytosis, membrane disruption, or passive diffusion.

Materials Magnetic Resonance Imaging of hESCs

H9 hESC (WiCell) hESC medium (see recipe) SPIO (Feridex, Bayer Healthcare Pharmaceuticals)



Protamine sulfate (PS, American Pharmaceutical Partners) Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS; see recipe) Heparin sodium (Fujisawa) Phantom (see recipe) Matrigel-coated 100-mm tissue culture dishes (BD Biosciences; coat dishes according to manufacturer’s instructions) 0.2-ml PCR tubes Signa 3T Excite HD scanner (GE Health System) and an array knee coil with GRE or SPGR sequence 1. Two days prior to the experiment, seed hESC onto Matrigel-coated 100-mm tissue culture dishes in 10 ml hESC medium. This procedure will allow removal of MEF and recovery of hESC culture.

2. One day prior to the experiment, dilute SPIO with hESC culture medium at a final concentration of 50 μg/ml. 3. Add clinical grade PS to the SPIO-containing medium to a final concentration of 6 μg/ml. Shake the mixture vigorously for 5 to 10 min. This procedure will enhance coating of SPIO with PS, generating SPIO-PS complex.

4. Incubate hESC with SPIO-containing medium overnight (8 to 12 hr) in the incubator. This incubation time varies depending on the cell types. We could achieve satisfactory labeling of hESC, human mesenchymal stem cells (hMSC), and mouse embryonic stem cells (mESC) after an 8-hr incubation.

5. Wash hESC two times, each time with 15 ml CMF-PBS. 6. Dilute heparin sodium with CMF-PBS to a final concentration of 10 U/ml. Wash hESC once with 15 ml heparin sodium–containing CMF-PBS. This procedure will allow elimination of SPIO-PS complex on the extracellular surface.

7. Suspend 2 × 106 hESC in 200 μl of CMF-PBS and transfer cell pellet to a 0.2-ml PCR tube. For the negative control, put the same number of non-labeled hESC in one PCR tube. The SPIO-labeled cell pellet will look dark-brown in color (Fig. 5A.3.3).

8. Place the PCR tubes onto the phantom gently to make sure no air is trapped in between the PCR tubes and the phantom to minimize any artifact from the air (Fig. 5A.3.3). The phantom will stabilize the PCR tubes and will prevent artifacts from the surrounding air. Air can induce hypointense signal on T2-weighted sequences. If a crack is noted on the phantom, it can be filled up with water to prevent artifacts.

9. Scan the hESC using T2-weighted sequences. We scan cells using Signa 3T Excite HD scanner and an array knee coil with GRE or SPGR sequence and the following parameters: TR (repetition time) = 100 to 200 msec, TE (echo time) = 20 to 30 msec, FOV (field of view) = 12 × 12 cm, and matrix = 192 × 192, NEX 1. MRI parameters should be optimized for different cell types, SPIO concentration, and magnetic field strength. For in vivo experiments, follow the same labeling method from steps 1 to 6 above. Then, transplant the optimal number of cells in the organ of interest. In our laboratory, 0.5 × 106 SPIO-labeled hESC are injected into the mouse heart for the myocardial infarction model.



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Figure 5A.3.3 Phantom for cellular imaging. The phantom is solidified within a plastic container. Any size plastic container can be used where PCR tubes can be placed stably. PCR tubes containing SPIO-labeled hESC cell pellets were placed onto the phantom. SPIO-labeled cells have a brown color (red arrows) and negative control, non-labeled cells look whitish (black arrow).

ALTERNATE PROTOCOL

DIRECT LABELING OF HUMAN EMBRYONIC STEM CELLS USING MANGANESE CHLORIDE SPIO has been employed to track and localize the transplanted stem cells with high sensitivity. However, this method does not monitor the viability of transplanted stem cells (Kraitchman et al., 2003). On the other hand, MnCl2 has been known to enter viable cells via voltage-gated calcium (Ca2+ ) channels. When the cells are biologically active, MnCl2 accumulates intracellularly to generate a T1-shortening effect to induce a hyperintense, bright signal on T1-weighted MRI acquisition (Aoki et al., 2006). The following protocol is for direct labeling of hESC using manganese and a T1-weighted MRI sequence.

Materials H9 hESC (WiCell) hESC medium (see recipe) TrypLE express (Invitrogen) MnCl2 (Sigma) 0.9% (w/v) sodium chloride (9 mg sodium chloride/ml of distilled water) Phantom (see recipe) Matrigel-coated 100-mm tissue culture dishes (BD Biosciences; coat dishes according to manufacturer’s instructions) Hemacytometer 15-ml conical tubes 0.2-ml PCR tubes Signa 3T Excite HD scanner (GE Health System) Magnetic Resonance Imaging of hESCs

1. Two days prior to the experiment, seed hESC onto Matrigel-coated 100-mm tissue culture dishes in 10 ml hESC medium.



2. On the day of experiment, remove the culture medium from the dish. Add 1.5 ml of TrypLE express and incubate the cells 5 min. Add 10 ml of culture medium and dissociate the cells by pipetting up and down several times. Wash the cells by centrifuging 5 min at 100 × g, room temperature. Resuspend the cells in culture medium and count cells using a hemacytometer. 3. Aliquot 3 × 106 hESC into one 15-ml conical tube. 4. Dissolve fresh MnCl2 with 0.9% sodium chloride solution to make a 0.1 mM MnCl2 solution. IMPORTANT NOTE: Always make a fresh MnCl2 solution. MnCl2 is easily degraded in solution.

5. Incubate the cells with 5 ml 0.1 mM MnCl2 for 30 min. Incubate negative control cells in 5 ml 0.9% sodium chloride alone. 6. Centrifuge the cells 5 min at 100 × g, room temperature. Aspirate the supernatant and wash the labeled cells twice, each time with 15 ml CMF-PBS. 7. Suspend each pellet in 200 μl CMF-PBS and transfer into 0.2-ml PCR tubes. Manganese-labeled pellets are usually white. For in vivo experiments, transplant these labeled cells in an organ of interest.

8. For an in vitro experiment, place the PCR tubes onto the phantom as described above (step 8 from the Basic Protocol). 9. Perform MRI scanning with T1-weighted spin echo sequence with the following parameters: TR; Repetition time = 800 msec. TE; Echo time = minimum, FOV; field of view = 12 × 12 cm; matrix = 192 × 192, NEX 1. MRI parameters should be optimized for the contrast effect depending on the cell type and magnetic field. Comparable magnetic labeling efficiency has been achieved with different hESC media such as DMEM/F-12 (Invitrogen) or mTeSR (STEMCELL Technologies).


Human embryonic stem cell medium (500 ml) 400 ml Knockout DMEM (Invitrogen) 100 ml Knockout serum replacement (Invitrogen) 5 ml of 10 mM non-essential amino acids (Invitrogen) 5 ml of 200 mM L-glutamine (Invitrogen) 3.5 μl of 14.3 M 2-mercaptoethanol (Sigma) 10 μg/ml recombinant human bFGF (R&D system) Store up to 1 week at 4◦ C Phantom 0.7% (w/v) agar (Sigma) 1% (w/v) copper sulfate (Sigma) Make solution with distilled water and microwave at medium high for 5 min Solidify in any plastic container before usage (Fig. 5A.3.3) Store up to 6 months at room temperature



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Phosphate-buffered saline, calcium- and magnesium-free (CMF-PBS) 0.144 g KH2 PO4 9.0 g NaCl 0.795 g Na2 HPO4 ·7H2 O H2 O to 1 liter Adjust to pH 7.4, if necessary Store indefinitely at room temperature COMMENTARY Background Information

Magnetic Resonance Imaging of hESCs

Magnetic resonance imaging (MRI) excites hydrogen protons, using a computerized program called the pulse sequence, to acquire and process the signal released from the excited protons. A pulse sequence consists of radiofrequency and gradient pulses which are carefully controlled in duration and timing to generate images of interest. In MRI, the critical properties are proton density and two basic relaxation times described as spin-lattice and spin-spin relaxation times denoted as T1 and T2, respectively. Relaxation time refers to the time required for the excited tissue to return to the equilibrium state after a radiofrequency pulse is applied. T1 and T2 depend on the proton density of each tissue. Fluid has longer T1 while fat has a shorter T1. T2 is usually shorter than T1 for a given tissue. Fluid has longer T2 while fat has a shorter T2. T2* refers to the effect of additional field inhomogeneity, which contributes to the dephasing signal. T2* is usually shorter than T2. In general, tissues with a long T2 give high signal intensities in T2-weighted images while a long T1 generates a weak signal. Exquisite intrinsic contrast achieved in MRI due to the differences in these relaxivity properties generates detailed images of the anatomy and morphology. Deeply located tissues in small animals can be visualized with high sensitivity by an MRI system. The location and number of the receive coil, the configuration of the coil element, and the magnetic field strength all play important roles. However, the ability of MRI to acquire images from any arbitrary tomographic plane enables detection of the cells in small animals with high sensitivity. In our laboratory, we image SPIO-labeled hESC in an 8-week-old SCID mouse using a 3.0 Tesla clinical scanner (Fig. 5A.3.4). Superparamagnetic iron oxide nanoparticles (size ∼100 nm) can induce a strong magnetic field inhomogeneity (dephasing signal) in the hydrogen atoms of water molecules during magnetic resonance imaging. When SPIO are taken up by the cells, the nanopar-

ticles create significant dephasing of protons, which consequently reduce T2* relaxation times. These properties enable robust visualization of SPIO-labeled cells through strong hypointense, negative signals described as the blooming effect (Bulte et al., 2001). Our data show significant contrast by in vitro MRI 14 days after labeling hESC with SPIO. In vivo MRI showed SPIO-induced contrast 20 days following transplantation of SPIOlabeled hESC in the mouse heart. Despite high sensitivity in the detection of the cells in the range of 10-9 mole/liter, this blooming effect may produce a large signal void at the region of interest to confound the MRI signal from the surrounding artifact and corrupt the anatomical details or the physiological function of the target tissue. Moreover, in vivo experiments have demonstrated that SPIO-labeled cells provide high sensitivity to detect the anatomical location of the cells. However, SPIO labeling does not provide any biologic information such as the viability of transplanted cells because of the non-specific uptake by the macrophages of the residual SPIO particles in the surrounding tissue from dead SPIO-labeled cells (Chen et al., 2008; Li et al., 2008). To address these limitations, our laboratory developed the Mn2+ labeling protocol for stem cells. Mn2+ is transported into the cellular cytoplasm of biologically active cells through a voltage-gated Ca2+ channel. These channels have high affinity for Ca2+ and its analog, such as Mn2+ , to accumulate within the cytoplasm by binding to specific sites on nucleic acid and intracellular proteins. Intracellular Mn2+ induces a T1-shortening effect, which allows clear delineation of the cells of interest with hyperintense, positive signal (Lin and Koretsky, 1997). Therefore, this contrast mechanism enables correlation between cellular viability and a T1-weighted positive signal. Our data shows Mn2+ -induced contrast effect is noted for 4 to 5 days after in vivo labeling (Fig. 5A.3.5). After these cells die, Mn2+ diffuses passively out of these dead cells. Consequently, decreased concentration of Mn2+



Figure 5A.3.4 In vivo MRI of hESC transplanted into the murine myocardium. Following direct labeling of hESC with SPIO and protamine sulfate, 0.5 × 106 hESC were transplanted into the left ventricle as indicated by negative dephasing signal (black arrow). This mouse was scanned using a 3T clinical MRI scanner (GE Healthcare System).

results in reduced T1-shortening effect and the contrast effect is lost. Currently, two distinct classes of iron oxide particles are available based on the hydrodynamic particle size. The mean diameter of SPIO is ∼100 nm. Ultrasmall superparamagnetic iron oxide particles (USPIO) are ∼40 to 50 nm. Both nanoparticles have similar chemical structures consisting of dextran coating to prevent destabilization and agglomeration of the colloidal suspension to enhance solubility in aqueous or biological media. Because of this chemical structure, both of these agents are biocompatible and SPIO is FDA-approved for imaging of liver lesions. Our studies have shown that mouse and human embryonic stem cell viability and differentiation capacity are not altered with SPIO labeling. However, other studies reported in vitro differentiation capacity of mesenchymal stem cells into chondrocyte lineage was reduced after SPIO labeling in a dose-dependent manner, whereas osteogenic and adipogenic differentiation was intact (Kostura et al., 2004). Direct SPIO labeling of hESC is simple and straightforward. Higher efficiency of iron-oxide labeling is achieved by adding

transfection agents such as PLL, PS or lipofectamine. All these transfection agents neutralize the negatively charged SPIO to facilitate the attraction and binding of slightly positive or neutral complex to the negatively charged cell membrane. The mechanisms by which these complexes enter the cell have not been completely investigated but they probably include endocytosis, invagination, or diffusion. Similarly, MnCl2 is a simple, robust, and safe method to label hESC. Mn2+ is an essential trace element in the human body with electrochemical properties analogous to Ca2+ . Using the voltage-gated Ca2+ channels, not only are the hESC localized with T1-weighted, positive signal, but the biological properties of the cells can also be determined. Nevertheless, the sensitivity/ability to detect cells is still higher with SPIO-labeled cells. However, drawbacks also exist for these contrast agents. First, the SPIO method requires long incubation times. To overcome this problem, advanced labeling methods such as magnetoelectroporation or magnetosonoporation have been reported (Walczak et al., 2005). In both methods, either a high-voltage electrical pulse or ultrasound was used to incorporate



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SPIO into the cytoplasm for a shorter time with a higher efficiency. Our data, however, demonstrated that electroporation alters the differentiation capability of mouse embryonic stem cells. In addition, the intracellular concentration of SPIO is diluted gradually with cell division or death. Second, at high doses of MnCl2, hESC toxicity occurs at the cellular level and systemic effects on neurologic and cardiovascular functions have been reported. These untoward effects of MnCl2 , however, have been overcome by concurrent calcium supplementation (Bruvold et al., 2005). The most successful optical method for imaging small numbers of transplanted cells in experimental models is bioluminescence. The sensitivity reaches 10−15 to 10−17 mole/liter enabling detection of 100 cells. This modality utilizes an internal biological light source, such as luciferase, which can be detected within the tissues of small animals using sensitive low-light imaging systems. Specific targeting of luciferase transgene expression in restricted cells and tissues of interest has allowed the localization and tracking of cell fate for studying a variety of disease processes. The CCD camera of the BLI is capable of detecting a minimum radiance of 100 photons per second per cm2 per steradian (photons/sec/cm2 /sr) and achieves a minimal image pixel resolution of 50 μm (Wu et al., 2003). High reproducibility (within ±8% SD from mean values) and detection sensitivity of this bioluminescence system for monitoring luciferase reporter gene expression has been demonstrated in vivo (Wu et al., 2001). While optical imaging detects signals from near-cellular level, this technique is limited to small animal imaging due to limited depth penetration of 1 to 2 cm, spatial resolution of 3 to 5 mm, and temporal resolution of

Magnetic Resonance Imaging of hESCs

seconds to minute (Auerbach et al., 1999). Clinical implementation of this technique (bioluminescence imaging) is not feasible.

Critical Parameters Although direct hESC labeling is simple and convenient, incubation time with SPIO and MnCl2 needs to be optimized. Satisfactory labeling of hESC could be achieved in an 8 to 12 hr incubation time with SPIO and in half an hour with MnCl2 without cytotoxicity. Excessive incubation time does not increase SPIO or MnCl2 uptake into cells but it does increases cytotoxicity.

Troubleshooting SPIO labeling is more sensitive than MnCl2 . With the imaging method described above, 50,000 hESC could be visualized with SPIO, while manganese chloride requires ∼106 hESC for direct MR visualization using a 3 Tesla clinical scanner. Care should be taken to make an MRI phantom as homogeneous as possible to remove any potential source of background artifacts such as air bubbles or cracks within the gelatin-based phantom.

Anticipated Results Hypointense, dark signals can be generated from the SPIO-labeled cells on T2*-weighted sequences (Fig. 5A.3.1). A visually distinct contrast can be observed starting at a magnetic field as low as 0.3T. Similarly, remarkable hyperintense, positive signals can be produced from the manganese-labeled hESC using T1weighted sequences (Fig. 5A.3.2). Significant bright signals can be achieved at a magnetic field as low as 1.5T.

Time Consideration

The entire procedure will take ∼72 hr from preparation of cells to MRI scanning of labeled

Figure 5A.3.5 In vivo manganese enhanced MRI of mESC transplanted into the murine right hind limb. The positive signal generated by mESC following intravenous administration of manganese is indicated by a black arrow. This mouse was scanned using a 3T clinical MRI scanner (GE Healthcare System).



cells. Labeling incubation time can be optimized from 4 to 12 hr with SPIO and from 30 min to 1 hr with MnCl2 . The MnCl2 solution must also be prepared on the day of cell labeling. Excessive incubation time may induce cytotoxicity.

Literature Cited Aoki, I., Takahashi, Y., Chuang, K.H., Silva, A.C., Igarashi, T., Tanaka, C., Childs, R.W., and Korefsky, A.P. 2006. Cell labeling for magnetic resonance imaging with the T1 agent manganese chloride. NMR Biomed. 19:50-59. Auerbach, M.A., Schoder, H., Hoh, C., Gambhir, S.S., Yaghoubi, S., Sayre, J.B., Silverman, D., Phelps, M.E., Schelbert, H.R., and Czernin, J. 1999. Prevalence of myocardial viability as detected by positron emission tomography in patients with ischemic cardiomyopathy. Circulation 99:2921-2926. Bruvold, M., Nordhoy, W., Anthonsen, H.W., Brurok, H., and Jynge, P. 2005. Manganesecalcium interactions with contrast media for cardiac magnetic resonance imaging: A study of manganese chloride supplemented with calcium gluconate in isolated Guinea pig hearts. Invest. Radiol. 40:117-125. Bulte, J.W., Douglas, T., Witwer, B., Zhang, S.C., Strable, E., Lewis, B.K., Zwicke, H., Miller, B., van Geleren, P., Moscovitz, B.M., Duncan, I.D., and Frank, J.A. 2001. Magnetodendrimers allow endosomal magnetic labeling and in vivo tracking of stem cells. Nat. Biotechnol. 19:11411147. Chen, I.Y., Greve, J.M., Gheysens, O., Willmann, J.K., Rodriguez-Porcel, M., Chu, P., Sheikh, A.Y., Faranesh, A.Z., Paulmurugen, R., Yang, P.C., Wu, J.C., and Gambhir, S.S. 2008. Comparison of optical bioluminescence reporter gene and superparamagnetic iron oxide MR contrast agent as cell markers for noninvasive imaging of cardiac cell transplantation. Mol. Imaging Biol. 11:178-187. Frank, J.A., Zywicke, H., Jordan, E.K., Mitchell, J., Lewis, B.K., Miller, B., Bryant, L.H. Jr., and Bulte, J.W. 2002. Magnetic intracellular label-

ing of mammalian cells by combining (FDAapproved) superaramagnetic iron oxide MR contrast agents and commonly used transfection agents. Acad. Radiol. 9:S484-S487. Kostura, L., Kraitchman, D.L., Mackay, E.M., Pittinger, M.F., and Bulte, J.W. 2004. Feridex labeling of mesenchymal stem cells inhibits chondrogenesis but not adipogenesis or osteogenesis. NMR Biomed. 17:513-517. Kraitchman, D.L., Heldman, A.W., Atalar, E., Amado, L.C., Martin, B.J., Pittenger, M.F., Hare, J.M., and Bulte, J.W. 2003. In vivo magnetic resonance imaging of mesenchymal stem cells in myocardial infarction. Circulation 107:2290-2293. Li, Z., Suzuki, Y., Huang, M., Cao, F., Xie, X., Connolly, A.J., Yang, P.C., and Wu, J.C. 2008. Comparison of reporter gene and iron particle labeling for tracking fate of human embryonic stem cells and differentiated endothelial cells in living subjects. Stem Cells 26:864-873. Lin, Y.J. and Koretsky, A.P. 1997. Manganese ion enhances T1-weighted MRI during brain activation: An approach to direct imaging of brain function. Magn. Reson. Med. 38:378-388. Suzuki, Y., Zhang, S., Kundu, P., Yeung, A.C., Robbins, R.C., and Yang, P.C. 2007. In vitro comparison of the biological effects of three transfection methods for magnetically labeling mouse embryonic stem cells with ferumoxides. Magn. Reson. Med. 57:1173-1179. Walczak, P., Kedziorek, D.A., Gilad, A.A., Lin, S., and Bulte, J.W. 2005. Instant MR labeling of stem cells using magnetoelectroporation. Magn. Reson. Med. 54:769-774. Wu, J.C., Sundaresan, G., Iyer, M., and Gambhir, S.S. 2001. Noninvasive optical imaging of firefly luciferase reporter gene expression in skeletal muscles of living mice. Mol. Ther. 4:297306. Wu, J.C., Chen, I.Y., Sundaresan, G., Min, J.J., De, A., Qiao, J.H., Fishbein, M.C., and Gambhir, S.S. 2003. Molecular imaging of cardiac cell transplantation in living animals using optical bioluminescence and positron emission tomography. Circulation 108:1302-1305.



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Lineage Tracing in the Intestinal Epithelium

UNIT 5A.4

Nick Barker1 and Hans Clevers1 1

Hubrecht Institute for Developmental Biology and Stem Cell Research, and University Medical Center Utrecht (UMCU), Utrecht, The Netherlands

ABSTRACT This unit describes the theory and detailed protocols for performing in vivo lineage tracing from Lgr5+ve intestinal stem cells using an Lgr5-EGFP-ires-CreERT2/Rosa26lacZ mouse model. Lineage tracing can be initiated in mice at any age by administering limiting doses of the hormone tamoxifen. This activates the lacZ reporter gene in the Lgr5+ve stem cells, which subsequently transmit this permanent genetic mark to their progeny as they repopulate the epithelium during normal homeostasis. Because the Lgr5+ve cells are long-lived, self-renewing stem cells, they continuously generate lacZ progeny, which contribute to tissue renewal over the entire lifetime of the mouse. The same protocols can be applied to performing in vivo lineage tracing from other Lgr5+ve stem cell populations, including those in the hair-follicle and stomach. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 13:5A.4.1-5A.4.11. Keywords: Lgr5 r in vivo lineage tracing r stem cell r intestine

INTRODUCTION The biology of the intestine is very well understood, yet the identity of the intestinal stem cells has remained elusive because of a lack of speciÞc markers (Barker et al., 2008). The authors recently identiÞed the Wnt target gene Lgr5 as a speciÞc marker for a restricted population of proliferating cells at the crypt base in both the small intestine and colon (Barker et al., 2007). In the small intestine, these Lgr5+ve cells are wedge-shaped cells called crypt base columnar cells (CBC), which are candidate stem cells found intermingled with the differentiated Paneth cells (Cheng and Leblond, 1974; Bjerknes and Cheng, 1981). To assess the stem cell potential of these Lgr5+ve cells, a mouse model was generated in which an EGFP-ires-CreERT2 expression cassette was inserted immediately downstream of the transcription start site of the endogenous Lgr5 promoter. The Lgr5+ve cells in this mouse consequently express an EGFP tag that allows visualization of them within the intestine using confocal microscopy and also allows us to efÞciently isolate them for in vitro analysis using FACS. The same Lgr5+ve cells also express a tamoxifen-inducible Cre (catalyzes recombination) enzyme that allows for in vivo lineage tracing when the mice are crossed with inducible reporter mice such as Rosa26lacZ (Soriano, 1999; Fig. 5A.4.1). Using this approach, it has been shown that a lacZ reporter gene activated in the Lgr5+ve cells is rapidly inherited by daughter cells, which constantly re-populate the intestinal epithelium as renewal occurs over the lifetime of the animal. In conclusion, this showed that the Lgr5+ve cells are the true stem cells of the intestine. Using a similar approach, it was also shown that Lgr5+ve populations in other tissues such as the hair-follicle (Jaks et al., 2008) and stomach (Barker et al., 2010) are adult stem cell populations responsible for maintaining tissue homeostasis. This unit provides step-by-step instructions for performing this in vivo lineage tracing using the Lgr5-EGFP-ires-CreERT2/Rosa26lacZ mouse model. The preparation and administration of the tamoxifen hormone (see Support Protocol) is described. There are

Current Protocols in Stem Cell Biology 5A.4.1-5A.4.11 Published online May 2010 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sc05a04s13 C 2010 John Wiley & Sons, Inc. Copyright



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Figure 5A.4.1 Image outlining in vivo lineage tracing using the Lgr5-EGFP-ires-CreERT2/Rosa26-lacZ mice. (A) Intestinal stem cells harboring the Lgr5-EGFP-ires-CreERT2 transgene express both EGFP and the CreERT2 enzyme. In the absence of tamoxifen, the CreERT2 enzyme is sequestered by heat-shock proteins in the cytoplasm. As a result, the lacZ reporter gene remains switched off due to the presence of a transcriptional roadblock. (B) Following induction, tamoxifen is taken up by the Lgr5+ve intestinal stem cells and complexes with the CreERT2 protein in the cytoplasm. This releases the CreERT2 enzyme from its heat-shock chaperones and allows it to enter the nucleus and catalyze the excision of the transcriptional roadblock from the lacZ reporter gene via Cre/loxP-mediated recombination. The lacZ reporter gene is consequently permanently switched on in the Lgr5-EGFP+ve stem cells. (C) These EGFP+ve /lacZ+ve stem cells subsequently divide to generate Lgr5-EGFP−ve progeny, which therefore inherit the genetic lacZ mark. These lacZ+ve progeny rapidly re-populate the intestinal epithelium, allowing their appearance and fate to be tracked over time.

protocols for optimal isolation and Þxation (see Basic Protocol 1) and lacZ staining of the intestines (see Basic Protocol 2). Finally, whole-mount (see Basic Protocol 3) and tissue section (see Basic Protocol 4) protocols for analyzing the lacZ staining are described.


NOTE: All protocols using live animals must Þrst be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow ofÞcially approved procedures for the care and use of laboratory animals.



ISOLATION AND FIXATION OF THE INTESTINE To visualize lineage tracing in the intestine, it is necessary to dissect, Þx, and lacZ stain the tissue from tamoxifen-treated Lgr5-EGFP-CreERT2/Rosa26lacZ mice.

BASIC PROTOCOL 1

Materials Tamoxifen-treated Lgr5-ires-CreERT2/Rosa26-lacZ mice (see Support Protocol) Gluteraldehyde lacZ Þxative (see recipe) or paraformaldehyde lacZ Þxative (see recipe) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 3-ml syringes and 21-G needles 50-ml centrifuge tube Rolling platform 1. Dissect the intestines from a tamoxifen-treated mouse and place into a petri dish containing 20 ml cold Þxative. 2. Equally divide the freshly isolated intestine into proximal, middle, distal, and colon segments and immediately ßush with 3 ml ice-cold gluteraldehyde Þxative to remove feces using a 3-ml syringe with a 21-G needle attached. An adult mouse small intestine is typically 40-cm long. The proximal segment is deÞned as the 12 to 14 cm immediately adjacent to the stomach. The distal segment is deÞned as the 12 to 14 cm immediately adjacent to the caecum. The intervening 12 to 14 cm deÞnes the middle segment. The colon is typically 7- to 8-cm long and is deÞned as the segment from the caecum to the rectum. The intestine is very susceptible to degradation following death because of its large microbial load. It is therefore crucial to isolate this tissue, ßush thoroughly, and initiate Þxation as soon as possible.

3. Place each intestinal segment into a separate 50-ml centrifuge tube containing 50 ml (∼20-fold excess) of ice-cold gluteraldehyde Þxative. Gluteraldehyde lacZ Þxative generally delivers optimal lacZ stains, but is incompatible with the majority of immunohistochemical procedures. PFA lacZ Þxative can be used in place of gluteraldehyde lacZ Þxative when lacZ/antibody co-stains are required.

4. Fix the intestines by constant mixing on a rolling platform for 2 hr at 4◦ C. Fixation times are critical—over-Þxation can destroy lacZ activity in the tissue.

5. Remove the Þxative and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform. Proceed to Basic Protocol 2.

TAMOXIFEN-INDUCTION OF IN VIVO LINEAGE TRACING In these studies, tamoxifen is used to induce the lineage tracing from the Lgr5+ve intestinal stem cells. The tamoxifen is prepared from a powder and injected into the Lgr5-EGFP/CreERT2/Rosa26lacZ mice.

SUPPORT PROTOCOL

Materials Tamoxifen powder (Sigma, cat. no. T-5648), stored at least 1 year at 4◦ C 100% ethanol Sunßower oil (supermarket variety) Adult Lgr5-iresCreERT2/Rosa26-lacZ mice (6 to 8 weeks old, ∼25 g; Jackson Laboratory) 37◦ C incubator 1-ml syringe and 25-G needle (BD Microlance)



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1. Dissolve the tamoxifen powder at 100 mg/ml in 100% ethanol by extensive vortexing at room temperature. 2. Add sunßower oil to achieve a 10 mg/ml tamoxifen/oil emulsion by extensive vortexing and store 0.4-ml aliquots up to 2 years at −20◦ C. 3. Pre-warm the tamoxifen stock (10 mg/ml) to 37◦ C and thoroughly vortex to ensure a homogenous emulsion. 4. Inject adult Lgr5-ires-CreERT2/Rosa26-lacZ mice intraperitoneally (i.p.) with 100 μl of 10 mg/ml tamoxifen (40 mg/kg) using a 1-ml syringe and 25-G needle. House induced mice under standard conditions. BASIC PROTOCOL 2

β-GALACTOSIDASE (lacZ) STAINING TO VISUALIZE INTESTINAL STEM CELLS The LacZ+ve progeny of the Lgr5+ve stem cells are visualized in the intestine by βgalactosidase staining.

Materials Fixed, freshly isolated intestines from tamoxifen-treated mice (see Basic Protocol 1) Equilibration buffer (see recipe) β-galactosidase (lacZ) substrate (see recipe) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 4% (w/v) paraformaldehyde (PFA; see recipe) Rolling platform 50-ml centrifuge tubes 1. Following the Þnal wash of the intestines (see Basic Protocol 1), remove the CMFPBS from the intestinal sections and add 50 ml equilibration buffer to each tube. Allow the intestines to equilibrate by constant mixing on a rolling platform 30 min at room temperature. 2. Transfer the intestines to a 50-ml centrifuge tube containing 50 ml of freshly-made lacZ substrate and allow the staining reaction to proceed with constant mixing on a rolling platform overnight at room temperature in the dark. X-gal (5-bromo-4-chloro-3-indolyl-β-galactosidase) is light-sensitive and incubation with this substrate and the subsequent post-Þxation in 4% PFA should therefore be performed in the dark.

3. Remove the staining solution and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform in the dark. 4. Remove CMF-PBS and add 50 ml (∼20-fold excess) of 4% PFA to each tube and Þx the intestines by constant mixing on a rolling platform overnight at 4◦ C in the dark. Optimal Þxation is achieved using a minimum 20-fold excess of 4% PFA at 4◦ C.

5. Remove the Þxative and wash the intestines two times for 10 min each in 50 ml CMF-PBS at room temperature on a rolling platform. Proceed to Basic Protocol 3. BASIC PROTOCOL 3 Lineage Tracing in the Intestinal Epithelium

WHOLE-MOUNT ANALYSIS OF lacZ STAINING IN THE INTESTINE Whole-mount segments of intestine are examined for the lacZ+ve progeny of the Lgr5+ve stem cells. The tissue is embedded in agarose, sectioned using a vibratome, and analyzed under a stereo microscope.



Materials Fixed and stained intestinal sections from tamoxifen-treated mice (see Basic Protocol 1) Phosphate-buffered saline lacking Ca2+ and Mg2+ (CMF-PBS) 4% low-melting-point agarose (Invitrogen, cat. no. 16520-100), pre-warmed to 40◦ C Glue (Bison, http://www.bison.nl, cat. no. Bi2058) Dissection pins Cardboard Petri dishes Stereo microscope (e.g., Olympus SZX9) linked to a digital camera Plastic basemolds (Klinipath, cat. no. 3051-P) Scalpel Vibratome (Microm model HM650V) Vibratome knives (Gillette, cat. no. 10) Starfrost microscope slides Coverslips (Menzel-Gläser) 1. To gain a global view of lacZ staining in the intestine, cut open a 1-cm piece from each intestinal segment along its length and pin it out onto a piece of cardboard with the inner surface (the surface epithelium) facing upwards. 2. Submerge the cardboard in a petri dish containing CMF-PBS and take whole-mount photos of the surface epithelium using a stereo microscope (e.g., Olympus SZX9) linked to a digital camera, with surface illumination (Fig. 5A.4.2). 3. To generate a more detailed overview of the lacZ staining present on local areas of surface epithelium, cut open a ∼1-cm piece of intestine along its length, lay it ßat in a tissue mold, and add 4% low-melting-point agarose until the tissue is completely submerged. 4. Allow the agarose to set (∼20 min), then remove the agarose/tissue block from the mold, and trim it to a perfect square using a scalpel blade. 5. Glue the block to the cutting platform of the vibratome so that the piece of intestine is perpendicular to the knife (i.e., side-on).

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Figure 5A.4.2 Whole-mount analysis of in vivo lineage tracing in the small intestine. (A) Lowpower image showing the presence of multiple lacZ+ve epithelial units throughout the small intestine 600 days post-induction. (B) LacZ+ve epithelial units visible on a 150-μm vibratome section of small intestine 5 days post-induction.



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6. Cut 150- to 200-μm sections (typically containing two to three crypts plus associated surface epithelium) and carefully transfer them to a microscope slide. 7. Place a coverslip over the sections to ensure that they remain ßat and take wholemount pictures as detailed above (Fig. 5A.4.2). BASIC PROTOCOL 4

DETAILED ANALYSIS OF lacZ STAINING ON TISSUE SECTIONS Tissue sections are prepared for closer analysis of lacZ expression.

Materials Intestinal tissue (see Basic Protocol 1) Tissue dehydration solutions: 70%, 80%, 96%, and 100% ethanol n-Butanol (Baker, cat. no. 8017) Liquid parafÞn (60◦ C) De-wax solvent (xylene; Klinipath, cat. no. 4055-9005) Tissue rehydration solutions: 100%, 96%, 90%, 80%, 70%, 60%, 50%, and 25% ethanol 0.1% (w/v) Neutral Red in ddH2 O Pertex mounting medium (Histolab) Tissue cassettes (Klinipath) Metal molds on an embedding station Heated forceps Cold plate (−12◦ C) Microtome 40◦ C water bath Starfrost microscope slides Hot-plate (∼55◦ C) Slide racks (Klinipath) Coverslips (Menzel-Gläser) Digital camera connected to a standard light microscope 1. Transfer the remaining intestinal tissues to a Klinipath tissue cassette and label the front panel clearly using a pencil. 2. Dehydrate the tissues by immersing the cassette in a 20-fold volume of 70% ethanol for 2 hr at 4◦ C. Refresh the 70% ethanol after 1 hr. Repeat this procedure using 96% ethanol and then 100% ethanol. Once tissues are transferred to 70% ethanol, they can be stored for up to 3 months at 4◦ C. The ethanol dehydration series cannot be interrupted after this stage. All dehydration steps using ethanol are performed at 4◦ C to ensure a gradual reduction in hydroxyl (water) bonds within the tissue, thereby reducing tissue damage.

3. Transfer the tissue cassettes to a 20-fold volume of n-butanol and incubate for 2 hr at room temperature. Following dehydration in the ethanol series, n-butanol must be used to remove the last traces of ethanol. Incubation in xylene will result in a loss of lacZ staining in the tissue.

4. Remove the tissue cassettes from the n-butanol and blot them onto tissues to remove any excess solvent. 5. Immerse the cassettes into 60◦ C liquid parafÞn overnight. Lineage Tracing in the Intestinal Epithelium

6. Remove the tissue cassettes from the liquid parafÞn and transfer them into metal molds on an embedding station. Carefully orient the tissues within the parafÞn using



heated forceps and then transfer the parafÞn blocks to a −12◦ C cold-plate for 30 min to allow them to solidify. 7. Prepare 6-μm thick sections using a microtome and transfer to a clean 40◦ C water bath. Allow the sections to spread out on the water and then ßoat them onto the upper surface of frosted microscope slides. 8. Dry the slides 1 hr on a 55◦ C hot-plate. 9. Transfer the slides into a slide rack and de-wax them two times by immersion in xylene, for 5 min each time. 10. Rehydrate the tissue sections by serial immersion in ethanol as follows:

1 min 1 min 1 min 1 min 1 min 1 min 1 min 1 min 1 min

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D

100% ethanol (two times) 96% ethanol 90% ethanol 80% ethanol 70% ethanol 60% ethanol 50% ethanol 25% ethanol ddH2 O.

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COLON DAY 1

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E

SI DAY 10

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Figure 5A.4.3 High-resolution examples of short-term and long-term lineage tracing in the small intestine and colon. (A) Isolated Lgr5-lacZ+ve stem cells present at the crypt base in the small intestine 1 day post-induction (black arrow). (B) Epithelial units in the small intestine partially populated by lacZ+ve progeny 5 days post-induction. LacZ+ve Paneth cells are typically not observed at these early time-points (red arrows). (C) Epithelial units in the small intestine entirely populated by lacZ+ve progeny 128 days post-induction. (D) Isolated Lgr5-lacZ+ve stem cells present at the colon crypt base 1 day post-induction (black arrow). (E) Epithelial units in the colon partially populated by lacZ+ve progeny 5 days post-induction. (F) Epithelial units in the colon entirely populated by lacZ+ve progeny 128 days post-induction.


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11. Counterstain by immersion in 0.1% Neutral Red solution for 1 min. Neutral Red is a counterstain that colors both the nucleus and cytoplasm red, generating an optimal contrast against the blue lacZ stain.

12. Quickly rinse the slides in 100% ethanol, three times for 30 sec each time, to remove excess Neutral Red counterstain and then transfer to xylene, incubate two times for 2 min each time. 13. Place a coverslip over the sections and seal it in place using Pertex mounting medium. 14. Photograph the sections using a digital camera connected to a standard light microscope (Fig. 5A.4.3).


β-galactosidase (lacZ) substrate 5 mM K3 Fe(CN)6 (see recipe) 5 mM K4 Fe(CN)6 ·3H2 O (see recipe) 2 mM MgCl2 (see recipe) 0.02% (v/v) NP40 (see recipe) 0.1% (v/v) sodium deoxycholate (see recipe) 1 mg/ml X-gal in CMF-PBS (see recipe) Prepare fresh and keep in the dark at room temperature Ethylene glycol tetraacetic acid Prepare 500 mM ethylene glycol tetraacetic acid (EGTA; Sigma) in ddH2 O. Adjust the pH to 7.2 using NaOH. Store indeÞnitely at room temperature.

Equilibration buffer 2 mM MgCl2 (see recipe) 0.02% (v/v) NP40 (see recipe) 0.01% (w/v) sodium deoxycholate in CMF-PBS (see recipe) Store indeÞnitely at room temperature Gluteraldehyde lacZ Þxative 1% (v/v) formaldehyde 0.2% (v/v) gluteraldehyde 0.02% (v/v) NP40 in CMF-PBS (see recipe) Prepare fresh and keep on ice Magnesium chloride Prepare a 1 M magnesium chloride (MgCl2 ) stock in CMF-PBS. Store indeÞnitely at room temperature.

NP40 Prepare a 10% (v/v) NP40 stock in CMF-PBS. Store indeÞnitely at room temperature.

Paraformaldehyde, 4% (w/v) Lineage Tracing in the Intestinal Epithelium

Dissolve 40 g paraformaldehyde (PFA; Sigma, cat. no. P6148) per liter of CMFPBS and heat to 60◦ C with constant stirring. Store up to 2 weeks (short-term) at 4◦ C or up to 6 months (long-term) at –20◦ C.



Paraformaldehyde lacZ Þxative 4% (w/v) paraformaldehyde (PFA; see recipe) 15 mM EGTA, pH 7.2 (see recipe) 2 mM MgCl2 in CMF-PBS (see recipe) Prepare fresh and keep on ice Potassium hexacyanoferrate II trihydrate (K4 Fe(CN)6 ·3H2 O) Prepare a 200 mM potassium hexacyanoferrate II trihydrate (K4 Fe(CN)6 ·3H2 O; Sigma, cat. no. P3289) stock in CMF-PBS. Store up to 2 weeks at 4◦ C.

Potassium hexacyanoferrate III (K3 Fe(CN)6 ) Prepare a 200 mM potassium hexacyanoferrate III (K3 Fe(CN)6 ; Sigma, cat. no. P8131) stock in CMF-PBS. Store up to 2 weeks at 4◦ C.

Sodium deoxycholate Prepare a 10% (w/v) sodium deoxycholate stock in ddH2 O. Store indeÞnitely at room temperature.

X-gal Prepare a 50 mg/ml X-gal (5-bromo-4-chloro-3-indolyl-β-galactosidase; Invitrogen, cat. no. 15520-018) stock in dimethylformamide (Sigma ACS grade 3, cat. no. 19937), dispense into 5-ml aliquots, and store up to 6 months at −20◦ C.

COMMENTARY Background Information The inner lining of the small intestine is arranged into multiple functional units of columnar epithelium comprising Þnger-like villi that protrude into the gut lumen, surrounded by ßask-shaped invaginations called crypts of Leiberkuhn (Stappenbeck et al., 2003; Sancho et al., 2004). These villi serve to maximize the surface area available for efÞcient absorption of digested food and water exiting the stomach. In the large intestine (colon), a ßat surface epithelium replaces these villi, reßecting its primary role in compacting undigested food remnants into stool/feces rather than absorption. The intestinal epithelium is probably the most rapidly renewing tissue in adults, undergoing a complete cycle of renewal every 5 days. This self-renewal is driven by a small population of self-renewing, multipotent stem cells located at the base of crypts (Bjerknes and Cheng, 1999). These stem cells generate a transient population of rapidly proliferating cells (the transit amplifying cells) that divide two to three times as they migrate upwards before differentiating into the major cell types present on the surface epithelium as they exit the crypts. These differentiated cells (comprising absorptive enterocytes, mucus-secreting goblet cells, and much rarer

hormone-secreting enteroendocrine cells) perform their essential functions as they continue migrating along the surface epithelium until Þnally dying by programmed cell death after 5 days. A fourth differentiated cell type in the small intestine called the Paneth cell escapes this upward migration and instead differentiates as it moves to the very base of the crypt. This cell lives up to 8 weeks and secretes antimicrobial lysozyme and cryptdins as part of the gut immune system. Lysozyme-secreting Paneth cells are not present in the colon, although cells with a similar function are thought to be present. In vivo lineage tracing in the Lgr5-iresCreERT2/Rosa26-lacZ mouse model centers on the inducible activation of a stablyintegrated lacZ reporter gene in the Lgr5+ve stem cell populations. Daughter cells derived from the lacZ+ve Lgr5 populations inherit this genetic lacZ mark, allowing their appearance and fate to be tracked in the corresponding tissue over time. This approach has been successfully used to demonstrate the stem cell function of Lgr5+ve cells in adult small intestine, colon, hair-follicle, and pyloric stomach. In the absence of tamoxifen induction, the Cre-ERT2 enzyme present exclusively in the Lgr5+ve stem cells is efÞciently sequestered by heat-shock proteins in the cytoplasm. The



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Rosa26lacZ reporter gene in the nucleus consequently remains switched off by virtue of the presence of a transcriptional roadblock sequence ßanked by loxP sites (Fig. 5A.4.1). Intraperitoneal injection of limiting doses of tamoxifen releases the Cre-ERT2 enzyme from its heat-shock protein chaperones and allows it to accumulate in the nucleus of isolated Lgr5+ve cells. Here, it rapidly mediates excision of the transcriptional roadblock by catalyzing recombination across the ßanking LoxP sites. The lacZ reporter gene is thereby activated in the Lgr5+ve cells and subsequently transmitted to their progeny as they repopulate the tissue during normal homeostasis (Barker and Clevers, 2007; Fig. 5A.4.1). Because the Lgr5+ve cells are long-lived, self-renewing stem cells, they continuously generate lacZ progeny, which contribute to tissue renewal over the entire lifetime of the mouse. LacZ reporter gene activity is visualized within the tissue using a 5-bromo-4-chloro3-indolyl-β-galactosidase substrate, which is catalyzed to a blue product by the lacZ enzyme.



In the Lgr5-EGFP-ires-CreERT2 mouse intestine, clusters of crypts silence the knock-in allele in a region-dependent fashion. The percentage of EGFP-ires-CreERT2+ve crypts consequently decreases from the proximal small intestine (∼70%) to the distal small intestine (∼30%). Such variegated expression of transgenes is commonly observed in the intestine. Importantly, however, no variegated expression is observed in other Lgr5+ve tissues, including the skin and stomach (nor is it observed in the intestine of an independent Lgr5-lacZ knockin model that the authors have used during the course of our studies). The Lgr5-EGFP-ires-CreERT2 mice are only viable as heterozygotes (i.e., one wildtype allele must always be present). In vivo lineage tracing is therefore always performed with Lgr5-EGFP-ires-CreERT2het /Rosa26lacZhet/hom mice. In principle, in vivo lineage tracing can be performed using Lgr5-EGFP-ires-CreERT2 mice in combination with any inducible reporter mouse strain that is activated using standard Cre/loxP technology. The dose of tamoxifen used to induce adult Lgr5-EGFP-ires-CreERT2/Rosa26-lacZ mice (estimated at 25 g total weight) is selected to achieve stochastic activation of the lacZ reporter gene within the Lgr5+ve population

of the intestine (to demonstrate that a single Lgr5+ve stem cell is responsible for driving the renewal of all cell types present on the crypt/villus epithelium). Higher doses of tamoxifen may be used to increase the frequency of lineage tracing if desired.

Anticipated Results

LacZ+ve Lgr5 stem cells at the crypt base should Þrst be observed 12 to 16 hr after induction. LacZ+ve Lgr5-derived transit amplifying cells will subsequently become visible in the crypts within 2 to 3 days. Expect to observe lacZ+ve progeny throughout the crypts and the associated surface epithelium 7 to 10 days postinduction. Typically, lacZ+ve enterocytes (alkaline phosphatase-positive) and goblet cells (PAS-positive) are present at this time-point because these exhibit the highest rate of turnover. Paneth cells (lysozyme-positive) have a much lower turnover rate and lacZ+ve examples are typically observed in the smallintestine only 3 to 4 weeks post-induction. At these early time-points, the surface epithelium in the small intestine contains both lacZ+ve and lacZ−ve progeny, creating a mosaic pattern (Fig. 5A.4.3B). This occurs because the surface epithelium is being supplied with cells from multiple crypts containing both lacZ+ve and lacZ−ve stem cells. After 60 to 80 days, the entire surface epithelium of tracing units is typically comprised of lacZ+ve cells, reßecting the fact that the lacZ+ve stem cell population has become dominant (with respect to the lacZ−ve stem cells) in the crypts supplying this intestinal unit. Given that the Lgr5+ve cells are long-lived, self-renewing stem cells, the frequency of lacZ+ve epithelial units present will remain constant even after the intestinal epithelium has undergone multiple rounds of complete renewal. Thus, expect to see multiple tracing units even 24 months after induction.

Time Considerations Following isolation of the intestines, the entire lacZ staining/Þxation/embedding procedure should take ∼4 days. Once embedded in parafÞn the tissues can be stored long-term at room temperature without loss of lacZ stain.

Literature Cited Barker, N. and Clevers, H. 2007. Tracking down the stem cells of the intestine: Strategies to identify adult stem cells. Gastroenterology 133:1755760.



Barker, N., van Es, J.H., Kuipers, J., Kujala, P., van den Born, M., Cozijnsen, M., Haegebarth, A., Korving, J., Begthel, H., Peters, P.J., and Clevers, H. 2007. IdentiÞcation of stem cells in small intestine and colon by marker gene Lgr5. Nature 449:1003-1007. Barker, N., van de Wetering, M., and Clevers, H. 2008. The intestinal stem cell. Genes Dev. 22:1856-1864. Barker, N., Huch, M., Kujala, P., van de Wetering, M., Snippert, H.J., van Es, J.H., Sato, T., Stange, D.E., Begthel, H., van den Born, M., Danenberg, E., van den Brink, S., Korving, J., Abo, A., Peters, P.J., Wright, N., Poulsom, R., and Clevers, H. 2010. Lgr5(+ve) stem cells drive self-renewal in the stomach and build longlived gastric units in vitro. Cell Stem Cell 6:2536. Bjerknes, M. and Cheng, H. 1981. The stem cell zone of the small intestinal epithelium III. Evidence from columnar, enteroendocrine, and mucosal cells in the adult mouse. Am. J. Anat. 160:77-91.

Cheng, H. and Leblond, C.P. 1974. Origin, differentiation, and renewal of the four epithelial cell types in the mouse small intestine. V. Unitarian theory of the origin of the four epithelial cell types. Am. J. Anat. 141:537-561. Jaks, V., Barker, N., Kasper, M., van Es, J.H., Snippert, H.J., Clevers, H., and Toftgard, R. 2008. Lgr5 marks cycling, yet long-lived, hair follicle stem cells. Nat. Genet. 40:12911299. Sancho, E., Batlle, E., and Clevers, H. 2004. Signaling pathways in intestinal development and cancer. Annu. Rev. Cell Dev. Biol. 20:695723. Soriano, P. 1999. Generalized lac-Z expression with the ROSA26 Cre expression strain. Nat. Genet. 21:70-71. Stappenbeck, T.S., Mills, J.C., and Gordon, J.I. 2003. Molecular features of adult mouse small intestinal epithelial progenitors. Proc. Natl. Acad. Sci. U.S.A. 100:1004-1009.

Bjerknes, M. and Cheng, H. 1999. Clonal analysis of intestinal epithelial progenitors. Gastroenterology 116:7-14.



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Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes in Mammalian Neocortical Neural Progenitors

UNIT 5A.5

Janice H. Imai,1,2 Xiaoqun Wang,1 and Song-Hai Shi1,2 1

Developmental Biology Program, Memorial Sloan-Kettering Cancer Center, New York, New York 2 BCMB Allied Program, Weill Cornell Medical College, New York, New York

ABSTRACT The importance of the centrosome in regulating basic cellular processes and cell fate decisions has become increasingly evident from recent studies tracing the etiology of developmental disorders to mutations in genes encoding centrosomal proteins. This unit details a protocol for a ßuorescence-based pulse labeling of centrioles of neural progenitor cells in the developing neocortex of mice. In utero electroporation of KaedeCentrin1 followed by in utero or ex vivo photoconversion allows a direct monitoring of the inheritance of centrosomes containing centrioles of different ages in dividing neocortical neural progenitors (i.e., radial glial cells). This is achieved by combining the irreversible photoconversion capacity of the Kaede protein from green to red ßuorescence with the faithful duplication of the centrosome during each cell cycle. After two mitotic divisions following photoconversion, mother centrosomes containing the original labeled centriole appear in both red and green ßuorescence, and can be distinguished from daughter centrosomes which appear in green ßuorescence only. This facilitates the study of the inheritance and behavior of the mother and daughter centrosomes in asymmetric cell divisions in the developing mammalian neocortex. Curr. Protoc. Stem Cell Biol. C 2010 by John Wiley & Sons, Inc. 15:5A.5.1-5A.5.12. Keywords: centrosome r Kaede-Centrin1 r mother and daughter centrosomes r photoconversion r neocortex r radial glia progenitor cell r in utero electroporation

INTRODUCTION This unit details a protocol for labeling the centrioles of neural progenitor cells (i.e., radial glia) in the developing neocortex of mice during the peak period of cortical neurogenesis (gestational days 13.5 through 17.5, i.e., E13.5 through 17.5), so as to study centrosome segregation in the context of neurogenesis (Wang et al., 2009). Centrioles/centrosomes of radial glia are initially labeled with green ßuorescence by in utero electroporation of a plasmid bearing the photoconvertible ßuorescent protein (Kaede) fused to the centriolar protein Centrin1 (Kaede-Centrin1) into the developing neocortex of embryos at E13.5. The plasmid is taken up by the radial glia within the ventricular zone. Approximately 24 hr later, each electroporated embryo is exposed to a brief pulse of violet light while in the uterus to convert the Kaede-Centrin1 protein from green to red ßuorescence. The uterus is placed back into the mouse, and the embryos are allowed to continue development. About 48 hr later, the brains of the embryos are recovered and two distinct populations of centrosomes can be observed: one has both green and red ßuorescence, representing the more mature mother centrosomes, and the other has green ßuorescence only, representing the less mature daughter centrosomes. We have shown that mother centrosomes are preferentially inherited by the renewing radial glia remaining in the ventricular zone, while daughter centrosomes are mostly inherited by the differentiating progeny that migrate away from the ventricular zone and occupy more Current Protocols in Stem Cell Biology 5A.5.1-5A.5.12 Published online October 2010 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470151808.sc05a05s15 C 2010 John Wiley & Sons, Inc. Copyright



dorsal layers of the neocortex, including the cortical plate (Wang et al., 2009). These results suggest that centrosome inheritance is tightly regulated and coordinated with cell fate decisions during asymmetric division of neural progenitors in the developing neocortex. This unit Þrst describes the well established method of in utero electroporation of plasmid DNA into radial glia in the developing neocortex (Basic Protocol 1). Next, a method for photoconverting Kaede-Centrin1 in vivo is described (Basic Protocol 2), followed by a procedure for preserving and visualizing the centrosomes with different ßuorescence spectra. Finally, an alternative procedure for photoconverting Kaede-Centrin1 in organotypic neocortical slices for time-lapse imaging studies of centrosome regulation during neurogenesis is presented (Alternate Protocol). NOTE: This protocol was developed in mice, and therefore some parameters must be determined empirically when applied to other species. NOTE: All solutions and equipment contacting the embryos should be sterile. NOTE: All protocols using live animals must Þrst be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow ofÞcially approved procedures for care and use of laboratory animals.

BASIC PROTOCOL 1

IN UTERO ELECTROPORATION OF THE KAEDE-CENTRIN1 PLASMID This protocol has been previously described (e.g., Tabata and Nakajima, 2008) and extensively employed in studying the function of gene(s) of interest in mammalian neocortical development.

Materials Timed-pregnant female mouse, E13.5 Isoßuorane Ethanol and iodine wipes Phosphate-buffered saline, sterile, 37◦ C Plasmid DNA (Kaede-Centrin1, 3.0 μg/μl; contact S.-H. Shi, [email protected]) mixed with Fast Green dye (Fisher Biotech) (1% w/v in PBS, 1 μl dye solution per 10 μl DNA solution); the use of endotoxin-free plasmid DNA (e.g., prepared using Qiagen Endotoxin-free Maxiprep kit) is recommended Antibiotic-PBS solution: penicillin (100 IU/ml)/streptomycin (100 mg/ml) in PBS, warmed to 37◦ C Antibiotic/analgesic solution (Duane Reade Triple Antibiotic Ointment plus Pain Relief; contains Bacitracin, zinc, neomycin sulfate, polymyxin B sulfate, and the analgesic pramoxine)

Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

Isoßurane induction chamber (VetEquip, e.g., cat. no. 901807) Isoßuorane dispenser (VetEquip) and nose cone Heating pad Disposable underpads Hair clipper Surgical instruments 10-ml sterile syringe Sterile gauze Sterile spatula Glass capillary injection needles (tip diameter ∼100 μm; beveled; see recipe) Electroporation system (BTX, ECM830, Harvard Apparatus)



Silk or nylon sutures Wound clips (7-mm; CellPoint ScientiÞc) Cotton-tipped applicators Surgically prepare the embryos 1. Anesthetize a timed-pregnant female mouse at 13.5 days of gestation (i.e., embryonic day 13.5 or E13.5) with isoßuorane in an induction chamber at a ßow rate of 1 to 4 liters/min (or according to the dispenser manufacturer’s recommendations). A single anesthetization procedure is carried out: Þrst, the animal is placed in the induction chamber until it is breathing but no longer moving; the air ßow valve from the isoßuorane dispenser to the chamber is closed while the air ßow valve from the dispenser to the nose cone is opened, and the animal is quickly transferred from the chamber to the nose cone (on the heating pad) while it is still unconscious.

2. Transfer the mouse from the isoßuorane induction chamber to the surgery table and onto a heating pad (37◦ to 40◦ C) covered with a clean disposable underpad. Place the mouse’s head in a nose cone connected to the isoßuorane output tube with the ßow rate of the isoßuorane set at 0.5 liter/min (or according to the manufacturer’s recommendations). 3. When the mouse is unresponsive to toe pinches, remove the abdominal fur with a hair clipper. Clean the shaven skin with iodine and alcohol wipes.

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Figure 5A.5.1 Procedure for in utero electroporation. (A) Timed-pregnant female mouse at gestational/embryonic day 13.5 (E13.5) under isoßuorane anesthesia with abdomen shaven and cleaned. The skin and underlying muscle have been cut open. (B) Mouse covered with a square of sterile gauze with a central hole cut out to expose only the opened abdominal cavity. The uterine horn containing embryos on the right side of the mouse has been gently lifted out of the cavity and placed on the gauze. (C) Positioning the embryo for injection by using a spatula to gently roll the embryo around within the yolk sac. (D) Glass capillary micropipet Þlled with plasmid DNA-dye mixture penetrating the lateral ventricle of the embryo through the uterine wall and yolk sac. (E) Positioning the electrodes for pulse delivery with the positive electrode covering the injected area and the negative electrode contacting the embryo at a diametrically opposed location. (F) Suturing the abdominal muscle after placing the uterine horn back into the abdominal cavity. (G) Applying wound clips to the skin.


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4. Make a vertical incision (∼2.5 cm) in the abdominal skin. To facilitate subsequent suturing of the muscle, separate the skin from the underlying muscle by cutting away the connective tissue. 5. Make a slightly smaller incision in the muscle (Fig. 5A.5.1A). From this point forward, the exposed abdominal tissue as well as the embryos should be continually moistened with warm (37◦ C) PBS dispensed dropwise from a 10-ml syringe.

6. Place a sterile gauze with a central hole cut out upon the abdomen such that only the incision area is exposed. 7. Pull the uterine horns gently out of the abdominal cavity and place on the gauze (Fig. 5A.5.1B).

Inject and electroporate the plasmid 8. Locate the lateral ventricles of the embryonic brain. At E13.5, the lateral ventricles of the developing brain occupy a large portion of each brain hemisphere and can be discerned through the uterine wall and yolk sac as a slightly darker, crescent-shaped area.

9. Gently rotate the embryo within its yolk sac with the aid of a spatula so as to position the head at an optimal angle for injection (Fig. 5A.5.1C). Take care not to squeeze the embryos. 10. Inject ∼1 μl plasmid DNA–dye mixture into the lateral ventricle of each embryo using a beveled glass micropipet (Fig. 5A.5.1D). As the DNA-dye mixture Þlls the ventricle, the ventricle will become more visible as a green crescent shape. In order to efÞciently perform the photoconversion (Basic Protocol 2) the following day and minimize surgery time, it is advisable to make the injection on the same side for each embryo. Only one ventricle per embryo is injected; there is no control per se.

11. After injection, pulse the embryo with a train of Þve 40- to 50-mV pulses (duration: 50 msec; interval: 950 msec) by covering the injected area with the positive electrode while the negative electrode maintains contact with the embryo on the diametrically opposite side of the head or body (Fig. 5A.5.1E). Radial glia line the ventricle in the area known as the ventricular zone; therefore, these take up the plasmid DNA when the voltage pulses are applied. As development proceeds, radial glia divide asymmetrically to produce radial glia that remain in the ventricular zone, as well as more fate-restricted daughter cells that migrate radially away from the ventricular zone to occupy more dorsal layers of the cortex. Care should be taken to avoid contact between the electrodes and the placenta during pulse application.

12. Repeat for each embryo. Generally, all embryos in a litter are injected.

Complete the surgery 13. Place the uterine horns back into the abdominal cavity and bathe in antibiotic-PBS solution. Suture and clip the wound (Fig. 5A.5.1F,G). Kaede-Centrin1 Labeling of Mother and Daughter Centrosomes

14. Apply analgesic ointment with a cotton-tipped applicator to the wound area. Place the mouse separately in a clean cage and closely monitor for respiratory distress until it is alert and ambulatory.



IN UTERO PHOTOCONVERSION OF THE KAEDE REPORTER Before undertaking this protocol, the minimum time required for complete photoconversion of Kaede-Centrin1 from green to red ßuorescence should be tested. For this calibration procedure, embryos are injected at E13.5, and the following day (E14.5) the embryos are exposed to violet light (see below); ∼3 to 5 min is usually sufÞcient. The light beam will penetrate the uterine wall, the yolk sac, and the outer tissues of the embryo’s head to the deepest layer of the developing neocortex, i.e., the ventricular zone where the Kaede-Centrin1-expressing radial glia are located. Embryos can be sacriÞced immediately following the photoconversion in these pilot experiments to examine the efÞcacy of the photoconversion.

BASIC PROTOCOL 2

Materials Pregnant mouse with electroporated embryos at E13.5 (Basic Protocol 1) Source of violet light (e.g., a ßuorescence dissection microscope with a mercury lamp and a 4 ,6-diamidino-2-phenylindole [DAPI] Þlter) Wound clip remover Additional reagents and equipment for anesthesia of the mouse and surgically removing and replacing embryos (Basic Protocol 1) 1. Approximately 24 hr after in utero electroporation, anesthetize the mouse with isoßuorane (see Basic Protocol 1) and reopen the wound by removing wound clips and sutures. 2. Expose the uterine horns and hold each embryo under the beam of a violet light (350 to 400 nm), which is focused on the injected area, for 3 to 5 min (Fig. 5A.5.2). 3. After all embryos are treated, place the uterine horns back into the abdominal cavity and resuture and clip the wound. 4. Apply appropriate analgesia and closely monitor the mouse until it has recovered, as described in Basic Protocol 1.

violet light beam

Figure 5A.5.2 Setup for in utero photoconversion. The embryo is held under a beam of violet light to effect photoconversion. The beam of light covers the injected area.



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ALTERNATE PROTOCOL

EX VIVO PHOTOCONVERSION IN BRAIN SLICES AND TIME-LAPSE IMAGING The same principles used for in vivo photoconversion can be applied to organotypic brain slice cultures, when the desired goal is a time-lapse imaging study of mother and daughter centrosome behavior over the course of neocortical neurogenesis. As for Basic Protocol 2, the minimum time required for complete photoconversion should be determined for slices in pilot experiments. Usually a much shorter time (18 kb in length can limit both the propagation of standard bacterial strains and the repertoire of unique restriction enzyme sites available to linearize the vector prior to electroporation.



pos se

lect .

3’

ar m

orter rep

ya

5’ h o

mo log y

log mo ho

*

*

neg . sele ct.

rm

vector backbone

Figure 5B.1.2 Plasmid map describing the possible structure of the targeting vector to generate a knock-in hESC line by homologous recombination. Two genomic fragments (5 homology arm and 3 homology arm) flank a reporter gene (green arrow) lacking an initiation codon, and a positive selection cassette (blue rectangle). The reporter gene is placed in frame with the ATG start codon included in the 5 homology arm. The red triangles represent loxP sequences that flank the positive selection cassette. A negative selection cassette (purple rectangle) can be included in the construct if desired. Unique restriction sites (marked as *) are located within the plasmid backbone to linearize the targeting construct prior to electroporation.

The homology arm sequences may also affect the success of homologous recombination. While targeting vectors containing nonisogenic sequences decrease the frequency of homologous recombination up to 20-fold in mESCs (te Riele et al., 1992; van Deursen and Wieringa, 1992), the efficiency of gene targeting in hESCs appears similar regardless of the origin of the homology arms. Practically, this also means that the same knock-in vector will target a given locus in different hESC lines at similar frequencies (Costa et al., 2007). Following recombination, positive selection cassettes are required to permit identification of stably transfected hESCs, which occur at a frequency between 1 in 104 105 electroporated cells. The neomycin phosphotransferase gene (neo) is highly expressed in hESCs when regulated by the mouse phosphoglycerate kinase (PGK) promoter (R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.), and this selection cassette is routinely used by the authors when transfecting hESCs. Additionally, if the hESCs are being cultured on mouse embryonic fibroblast feeder cells, geneticin (G418)-resistant mouse lines are readily available. As an alternative, the neo gene may also be replaced with the hygromycin B phosphotransferase (hph) gene and the antibiotic hygromycin B used to enrich for stably transfected hESCs (L. Azzola, A.G. Elefanty, and E.G. Stanley, unpub. observ.). In correctly targeted clones, retention of the positive selection cassette in the genome can influence expression of the target locus and of neighboring genes (Hug et al., 1996; Pham et al., 1996; Scacheri et al., 2001; R. Davis, A.G. Elefanty, and E.G. Stanley, unpublished observations). Therefore

Generic Manipulation of Stem Cells


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flanking the cassette with loxP sequences allows for subsequent removal by transient expression of Cre recombinase (Gu et al., 1993). Replacement-type targeting vectors can also include a negative selection cassette, such as the herpes simplex virus thymidine kinase (HSVTK) gene, to allow for selection against random integrants and enrich for targeted recombinants (Zwaka and Thomson, 2003). This cassette is located outside the region of homology to the target gene, normally at the end of the short homology arm.

Culturing the hESCs The adaptation of the hESCs to enzymatic passaging using either trypsin or TrypLE Select (Invitrogen) is essential for success in the protocols described. Without such preconditioning, disaggregation of hESCs to a single-cell suspension results in extensive cell death (UNIT 1C.1). Protocols for the adaptation and expansion of enzymatically passaged hESCs are described in detail in UNIT 1C.1 and UNIT 1D.3. Only hESCs that have undergone 5 to 10 enzymatic passages should be used in Basic Protocols 1 and 2, and Support Protocol 4. Validation of hESC Reporter Knock-in Lines The protocols in this unit describe the generation of hESC reporter lines by homologous recombination, identification of correctly targeted clones, removal of the selectable marker, and single-cell cloning of the cell line. However, before any hESC reporter line is used for experimentation, further screening procedures are required to validate the cell line. Southern blot analysis of the putatively targeted hESC clones can confirm that the desired recombination events have occurred at both the 5 and 3 ends, and that the cells contain only a single copy of the reporter gene. A detailed description of Southern blotting is outside the scope of this unit. In brief, genomic DNA extracted from the hESC colonies is digested with restriction enzymes, electrophoresed on an agarose gel, and transferred to a membrane. This membrane is hybridized with radiolabeled probes to the gene locus that lies external to the region of homology with the targeting vector, and the labeled DNA fragments detected using either a phosphoimager or X-ray film. Such probes serve to identify correctly targeted alleles by virtue of size differences between hybridizing fragments generated by wild-type and targeted alleles. These size differences arise because of the incorporation of additional DNA sequences and/or new restriction enzyme sites associated with the introduction of the reporter gene and selectable marker sequences into the native locus. In addition, a radiolabeled probe that hybridizes with the coding sequence of the reporter gene can confirm that only a single integration event occurred. The genetically manipulated hESC line must also maintain the characteristics of a stem cell. The cells should be regularly analyzed by flow cytometry for the expression of a panel of stem cell markers including transcription factors, such as OCT4 and NANOG, and the cell surface antigens, such as SSEA3 and 4, Tra-1-60, Tra-1-81, and CD9 (UNIT 1B.3). Additionally, when the genetically modified hESCs are injected into the testes of immunodeficient mice, they should form multi-lineage teratomas (UNIT 1B.4). The karyotype should also be periodically checked to confirm that no karyotypic abnormalities have been introduced during the genetic manipulations. It is recommended that this analysis, to at least the level of G-banding, be performed by a clinical cytogenetics facility. Generation of hESC Reporter Knock-In Lines

NOTE: The following protocols are performed in either a Class I (laminar-flow) biosafety cabinet or a Class II biohazard hood.



NOTE: All materials and reagents that come into contact with live cells must be sterile and proper aseptic technique must be used when handling the cells or setting up experiments. NOTE: All incubations are performed in a 37◦ C, 5% CO2 humidified incubator, unless otherwise specified.

ELECTROPORATION OF hESCs AND SELECTION OF ANTIBIOTIC-RESISTANT hESCs

BASIC PROTOCOL 1

Electroporation is the most common means of generating targeted mESC lines (Giudice and Trounson, 2008). While the number of reports describing gene targeting by homologous recombination in hESCs is currently limited, the majority of these have also utilized electroporation (Zwaka and Thomson, 2003; Costa et al., 2007; Irion et al., 2007; Davis et al., 2008a; Braam et al., 2008). This protocol describes the procedure for electroporating hESCs with a linearized gene-targeting vector, followed by the positive selection of stably transfected colonies. It is recommended that the neo gene be included in the positive selection cassette, allowing for the drug geneticin (G418) to be used as the selection agent.

Materials hESCs in 150-cm2 tissue culture flasks at enzymatic passage 5 to 10 (see UNIT 1C.1) in hESC medium (see recipe) 150-cm2 gelatinized tissue culture flasks (see recipe) preseeded with mitotically inactivated MEFs at 1 × 104 /cm2 for passaging hESCs prior to electroporation MEF medium (see recipe) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) hESC medium (see recipe), 37◦ C Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) 0.4% (w/v) Trypan blue (Fluka) Soybean trypsin inhibitor (see recipe; Invitrogen), optional Linearized targeting vector (see Strategic Planning) in Tris/EDTA (TE) buffer (see recipe) for transfection 60-mm gelatinized tissue culture dishes preseeded with 2 × 104 /cm2 mitotically inactivated MEFs Geneticin/G418 Selective Antibiotic (Invitrogen) Mitotically inactivated, irradiation-treated (UNIT 1C.3) G418-resistant mouse embryonic fibroblasts (MEFs; Conner, 2000) 37◦ C water bath Gene Pulser cuvette, 0.4-cm electrode gap, sterile (Bio-Rad, cat. no. 165-2088) 15- and 50-ml sterile centrifuge tubes Refrigerated centrifuge Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Tissue culture microscope with phase contrast objectives and phase rings Hemacytometer (Neubauer) Electroporator (Gene Pulser II System; Bio-Rad) Sterile Pasteur pipets Prepare hESCs prior to electroporation (day −1, day 0) 1. On the day prior to electroporation, enzymatically dissociate the hESCs from two 150-cm2 flasks and replate onto two fresh 150-cm2 flasks preseeded with MEFs at low density (1 × 104 MEFs/cm2 in 20 ml MEF medium), using either 2 ml trypsin or 2 ml TrypLE Select per 150-cm2 flask (Fig. 5B.1.3A). It is critical that hESCs are adapted to enzymatic passaging using either trypsin or TrypLE Select prior to beginning this protocol. The enzymatic expansion of hESCs from



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A

B

C

D

Figure 5B.1.3 Photomicrographs of hESCs expanded in bulk culture, after electroporation, during drug selection, and following selection. Some groups of hESCs or individual colonies in panels (A), (B), and (C) are outlined by black-dotted lines. (A) hESCs in bulk culture grown on MEFs at reduced density on day of electroporation. (B) hESCs on the day following electroporation. (C) hESCs 5 days after electroporation, just prior to antibiotic selection. (D) A hESC colony following 5 days of selection. (A) through (D) Original magnification 50×.

stock cultures maintained by mechanical passaging is described in detail in UNIT 1C.1. The cells should be enzymatically disaggregated every 3 or 4 days, and split no more than at a ratio of 1:2. Generally, after the first 4 or 5 enzymatic passages, the hESCs will have been expanded into two 150-cm2 flasks. The hESCs are passaged onto fresh MEFs at a ratio of 1:1 on the day prior to the electroporation to ensure that the cells are actively dividing and to remove dead and dying cells. Seeding the flasks with a reduced density of MEFs (1 × 104 MEFs/cm2 compared with 2 × 104 MEFs/cm2 ), results in a partial depletion of feeder cells.

2. At a time point ∼2 to 3 hr before the electroporation, aspirate the medium and re-feed the cells with fresh hESC medium. 3. At a time point ∼2 hr before the electroporation, place a 10-ml aliquot of CMF-PBS on ice and a 40-ml aliquot of hESC medium in a water bath at 37◦ C. 4. Place the Gene Pulser electroporation cuvette on ice.

Harvest the cells 5. Harvest the hESCs passaged the day before. First, aspirate the hESC medium and then rinse the flasks with 5 to 10 ml CMF-PBS. 6. Add 2 ml of trypsin or TrypLE Select to each 150-cm2 flask and ensure that the dissociation solution coats the surface of the cells. Incubate 4 min at 37◦ C in the incubator. Check that the hESCs dislodge from the flasks with gentle tapping. Generation of hESC Reporter Knock-In Lines

If the cells are dissociated with trypsin, then a neutralization step is required. Add 1 ml of soybean trypsin inhibitor to each flask and swirl to mix. A specific neutralizing agent is not required for TrypLE Select.



7. Add 8 ml of hESC medium to each flask, mix, and transfer the contents of the two flasks to a single 50-ml centrifuge tube. 8. Pellet the cells by centrifuging the tube 3 min at 480 × g, at 4◦ C, and aspirate the supernatant. 9. Resuspend the hESCs in 10 ml of CMF-PBS and transfer the cell suspension to a 15-ml centrifuge tube. 10. Using a Gilson pipettor, mix 10 μl of the cell suspension with 10 μl trypan blue. Load 10 μl onto a hemacytometer and, using a tissue culture microscope, perform a cell count (UNIT 1C.3). Subtract the total MEF number (3 × 106 ) from the count. A total of 1 × 107 hESCs is required per electroporation. Two 150-cm2 flasks seeded with MEFs at a density of 1 × 104 /cm2 will contain ∼3 × 106 MEFs. After subtracting the MEF count, a semi-confluent 150-cm2 flask of hESCs typically contains between 6–8 × 106 hESCs. Therefore the yield from two such flasks (∼12–16 × 106 hESCs) will provide enough hESCs for a single electroporation (∼10 × 106 hESCs). The remaining hESCs can be replated onto a new flask seeded with MEFs at the regular density (2 × 106 MEFs/cm2 ) to maintain the undifferentiated culture. If there are ∼2–4 × 106 hESCs remaining, these should be plated onto a 75-cm2 flask, while a 150-cm2 flask should be used if there are in excess of 4 × 106 hESCs.

Prepare the cells with the targeting vector for electroporation 11. Centrifuge the cells again for 3 min at 480 × g, 4◦ C, aspirate the supernatant, and resuspend 1 × 107 hESCs in a final volume of 750 μl ice-cold CMF-PBS. 12. Using a Gilson pipettor, add 50 μl of TE buffer containing between 10 and 20 μg of the linearized gene-targeting vector to the electroporation cuvette. 13. Carefully transfer the hESC suspension into the cuvette using a pipettor, ensuring that the DNA-containing TE buffer and hESCs are evenly resuspended. 14. Place the cuvette containing the DNA and cell suspension mix on ice for 5 min.

Electroporate the cells 15. Wipe the outside of the cuvette to remove any water or ice before electroporating the cells at 250 V and 500 μF (Costa et al., 2007). Other groups have achieved successful transfection of the plasmid by electroporating the hESCs at 320 V and 200 μF (Zwaka and Thomson, 2003), or 320 V and 250 μF (Braam et al., 2008). In our laboratory, our conditions have been used to target at least 9 loci (Costa et al., 2007; A.G. Elefanty and E.G. Stanley, unpub. observ.).

Plate the electroporated cells 16. Using a Pasteur pipet, transfer the contents of the cuvette to a 50-ml centrifuge tube containing 10 ml of prewarmed hESC medium. 17. Centrifuge the electroporated cells 3 min at 480 × g, room temperature (20◦ to 25◦ C). 18. Carefully aspirate the supernatant and gently resuspend the pellet in 15 to 18 ml of prewarmed hESC medium. Steps 16 to 18 remove cellular debris that could impair the viability of the surviving hESCs.

19. Plate the cell suspension into five or six 60-mm dishes preseeded with 2 × 104 MEFs/cm2 , and incubate the dishes in a humidified incubator at 37◦ C, 5% CO2 (Fig. 5B.1.3B).



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The number of 60-mm dishes that the electroporated hESCs are plated into is dependent on the proportion of hESCs that survive the electroporation procedure. This percentage not only varies between hESC lines, but also reflects the degree to which the cells have adapted to the enzymatic passaging prior to electroporation. Plating the hESCs into five or six 60-mm dishes usually ensures the hESC density is low enough that the cells can be left to recover and proliferate for 4 to 5 days before beginning selection. If the hESC cultures are confluent in less than 4 days, plate the cells into a larger number of 60-mm dishes when performing future electroporations.

20. Two days after the electroporation, gently aspirate the medium containing the dead cells from each dish and replace with 4 ml of fresh hESC medium per dish. Repeat this daily.

Select for geneticin-resistant hESCs (day 4 or 5) 21. Supplement 200 ml of hESC medium with the drug geneticin (G418) so that the final concentration is 50 μg/ml. The recommended final concentration of G418 to use is 50 μg/ml; however, the optimal concentration for selection may vary for different hESC lines. Prior to performing an electroporation, determine the minimum concentration of G418 required to eliminate G418-sensitive hESCs within 5 days of addition. For 1 week of selection, 200 ml of hESC medium containing G418 should be sufficient stock. This medium has a limited lifespan of 7 days.

22. Apply 50 μg/ml G418 selection to the hESCs when the dishes are ∼60% to 80% confluent with hESCs (4 or 5 days following electroporation; Fig. 5B.1.3C). A 60-mm dish that is semi-confluent with nontransfected hESCs should also be placed under selection as a control.

23. Change the selection medium daily. Frequent medium changes are necessary during selection to remove the dead cells.

24. Four days into selection, supplement the dishes with fresh MEFs at 1 × 104 MEFs/cm2 . Resuspend the required number of MEFs in 20 ml hESC medium containing G418 and dispense evenly over all the dishes, such that each dish contains a total volume of 4 ml. MEFs can support hESC growth for ∼1 week, and so the dishes will need to be supplemented with additional MEFs during the protocol to maintain densities of ∼2 × 104 MEFs/cm2 . This should be done either 8 days after the initial plating of the electroporated hESCs (Fig. 5B.1.3D), or if the MEF density appears low after the onset of selection. Supplementation with fresh MEFs is required even if the MEFs are G418 resistant.

Remove the selection agent 25. Following 7 days of G418 selection, return to culturing the G418-resistant cells in hESC medium without G418. After 7 days, the control 60-mm dish should contain no residual viable hESCs. If live hESCs remain on the control dish, continue G418 selection on all dishes until colony transfer (∼18 days post electroporation). Alternatively, the dose of G418 required to kill control hESCs may need to be re-titrated.

26. Allow the colonies to grow for approximately a further 7 days, changing the hESC medium daily. The colonies are ready to transfer to 48-well tissue culture plates (Support Protocol 1) when they are ∼2 mm in diameter. If the cells begin to differentiate, they should be transferred sooner. Generation of hESC Reporter Knock-In Lines



PICKING AND EXPANDING ANTIBIOTIC-RESISTANT hESC COLONIES This protocol describes transferring and expanding individual genetically modified hESC colonies that emerge on the 60-mm dishes following the selection procedures described in Basic Protocols 1 and 2. Approximately 7 to 9 days after the withdrawal of the selection agent from the culture medium, each hESC colony is mechanically passaged into a well of a 48-well tissue culture plate (Fig. 5B.1.4A,B). In our experience, the number of antibiotic-resistant colonies after Basic Protocol 1 may vary from 50 to more than 300 per 107 electroporated hESCs. One week after the initial transfer, the hESCcontaining plates are then duplicated, with one plate destined for extraction of DNA to perform the PCR screen (Support Protocols 2 and 3), while the other plate will serve to maintain the hESC colonies in culture until positive clones are identified.

SUPPORT PROTOCOL 1

Materials Flat-bottomed 48-well tissue culture plates, gelatinized (see recipe) and preseeded with mitotically inactivated MEFs at 2 × 104 /cm2 Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) hESC medium (see recipe) 60-mm tissue culture dishes containing drug-resistant hESC colonies from either Basic Protocol 1 or 2 26-G, 1/2 -in. (0.45 × 13–mm) needles 1-ml syringe 200-μl pipet tips Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Stereomicroscope Prepare 48-well tissue culture plates for transfer of the hESC colonies (day −1) 1. Estimate the total number of individual, undifferentiated hESC colonies to be picked from the tissue culture dishes. Seed enough 48-well plates with ∼0.75 × 106 (MEFs in 12 ml MEF medium, 250 μl/well) MEFs/plate, such that each hESC colony can be transferred to an individual well. These 48-well plates are designated the “Primary” plates.

Figure 5B.1.4 Photomicrographs of a hESC colony being transferred and expanded. (A) 25× magnification of a hESC colony sliced into a grid pattern prior to dislodgement and transfer to a well on a 48-well plate. (B) Transferred pieces from a single hESC colony grown for 7 days in a well on a 48-well plate.



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It is recommended that one person should not pick more than 150 hESC colonies at a time due to the difficulty in maintaining and processing large numbers of colonies later in the PCR screening (Support Protocol 3). If the electroporation performed in Basic Protocol 1 has been particularly successful, a pair of researchers can manage to pick, replicate, and process a larger number of antibiotic-resistant colonies.

Pick hESC colonies (day 0) 2. Aspirate the MEF medium from the 48-well plates pre-seeded with MEFs, and dispense 200 μl fresh hESC medium into each well. 3. Aspirate the hESC medium from the 60-mm dishes containing the hESC colonies and supplement with 4 ml of fresh hESC medium. 4. Using a 26-G needle attached to a 1-ml syringe, cut a hESC colony in a grid motif to generate a minimum of 16 small pieces (Fig. 5B.1.4A). Detach the pieces from the dish by flicking them off with the needle, or with a 200-μl pipet tip attached to a pipettor. These procedures are performed under a zoom-focus stereomicroscope.

5. Using a 200-μl pipet, collect all the pieces and transfer them into a well on one of the “Primary” 48-well plates. 6. Discard the 26-G needle and 200-μl pipet tip after harvesting the hESC colony. 7. Repeat steps 4 to 6 to harvest the remaining hESC colonies on the tissue culture dishes. Select well-spaced colonies to avoid cross-contamination of clones. Avoid picking colonies that are significantly smaller than the majority of the colonies. Smaller colonies can be left to expand longer on the 60-mm plates and picked several days later when larger.

8. Replace the hESC medium in the wells daily with 200 μl of fresh medium per well. Within 2 days of picking, the hESCs should be visible with the formation of multiple colonies in each well.

Replicate the hESC-containing 48-well plates (days 5 to 7) 9. Between 5 and 7 days after picking the hESC colonies, verify that the wells are approaching confluence (Fig. 5B.1.4B). Prepare two sets of 48-well plates with MEFs (as described in step 1). Label one set of plates “DNA,” and the other set “Maintenance.” 10. The next day, replace the MEF medium on the multi-well plates labeled “DNA” and “Maintenance” with 100 μl of fresh hESC medium. 11. Replace the hESC medium on the 48-well plates labeled “Primary” with 300 μl of hESC medium. 12. Using a plugged 200-μl pipet tip attached to a pipettor, scrape the bottom of a well on a “Primary” plate in a criss-cross fashion until the hESC colonies have detached. Examine the well under a stereomicroscope to confirm that the hESC colonies have broken into small cell clumps. If the hESCs have not fragmented into small clumps, pipet the cells up and down to try to break up the colonies.

Generation of hESC Reporter Knock-In Lines

13. Transfer 100 μl of the well contents to a 48-well plate labeled “Maintenance,” and the remaining 200 μl to the corresponding well on the plate labeled “DNA.” Expel air bubbles into the medium to assist in identifying wells that contain passaged hESCs. Discard the 200-μl pipet tip after use.



Uneven distribution of the hESC clones between the 48-well plates allows the plates labeled “DNA” to be processed and screened by PCR before the hESCs on the “Maintenance” plates have reached confluence and need to be passaged. If the hESCs in the wells of the “Primary” plate are confluent, and if there are fewer than 48 hESC clones to screen, it is often convenient to directly isolate DNA for PCR screening at this stage without further expanding the cell numbers. Instead of seeding the cells into the wells of the “DNA” plate, transfer the hESC clones into individually labeled microcentrifuge tubes, and pellet the cells by centrifuging 3 min at 480 × g, 4◦ C. The genomic DNA can be isolated using the method described in the Alternate Protocol.

14. Repeat steps 12 and 13 to transfer all remaining hESC colonies. 15. The next day, replace the medium on all plates with 200 μl of fresh hESC medium. Change the hESC medium daily. 16. Approximately 6 days after duplicating the hESC clones, the hESCs on the “DNA”labeled plates will be confluent enough to isolate sufficient DNA for PCR screening (Fig. 5B.1.4B). Proceed to Support Protocol 2 for instructions on how to process these plates. 17. While screening the hESCs to identify clones that contain the correct genetic modification, continue to change the hESC medium daily on the “Maintenance” plates. If the majority of hESC-containing wells on the “Maintenance” plates are more than 80% confluent, or the hESC colonies have begun to differentiate before the PCR screening is complete, the cells will need to be passaged. This is performed as described in steps 9 to 14, but not in duplicate.

PREPARATION OF GENOMIC DNA FROM hESCs GROWING IN 48-WELL TISSUE CULTURE PLATES

SUPPORT PROTOCOL 2

To maximize the quantity of genomic DNA obtained from the genetically modified hESC clones, the cells can be let to overgrow and the plates should only be harvested when the majority of the clones are confluent. This protocol describes a simple, but effective, method of DNA extraction from hESCs growing in 48-well plates. This DNA is then used in a PCR screen to identify hESC colonies that contain the correct genetic modification. In some cases, the PCR screen is not sensitive enough to identify positive clones from DNA isolated using this method. Under these circumstances, the DNA should be extracted from the hESCs incorporating a phenol/chloroform extraction step as described in the Alternate Protocol.

Materials 48-well tissue culture plates labeled “DNA,” containing confluent hESC colonies (from Support Protocol 1, step 13) Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) DNA lysis buffer containing 200 μg/ml proteinase K (see recipe) 100% (v/v) and 70% (v/v) ethanol Tris/EDTA buffer (TE buffer; see recipe) Temperature-adjustable incubator Centrifuge with multi-well plate spinner attachment Blotting paper Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips 1. Aspirate the medium from the plates assigned for DNA extraction and rinse the wells with 200 μl CMF-PBS.



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2. Add 100 μl DNA lysis buffer containing 200 μg/ml proteinase K to each well and incubate 3 hr at 55◦ C. Approximately 5 ml of lysis buffer is required for each plate. Alternatively, the plates can be incubated overnight at 37◦ C.

3. Add 250 μl of 100% ethanol to each well and mix by gently tapping the plate. Mix the solutions until a DNA precipitate is visible in each well.

4. Centrifuge the plates 15 min at 3000 × g for 15 min, room temperature, to pellet the DNA in the bottom of each well. 5. Decant the supernatant by carefully inverting the plate on blotting paper. Take care to ensure that the DNA precipitates remain in the wells.

6. Add 250 μl of 70% ethanol to each well. Centrifuge the plates again 15 min at 3000 × g, room temperature, to ensure the DNA is pelleted on the bottom of the wells. This step removes residual salt from the DNA pellet.

7. Remove the 70% ethanol solution from each well. First, invert the plate on blotting paper and then carefully remove any residual ethanol in each well using a 200-μl pipet with a plugged tip attached. Leave the DNA to air dry for no more than 10 min at room temperature. Use a different 200-μl plugged tip for each well.

8. Resuspend the DNA in 100 μl of TE buffer and incubate for 3 to 4 hr at 55◦ C to dissolve the pellet. The DNA can be stored for at least 3 months at 4◦ C. ALTERNATE PROTOCOL

ISOLATION OF GENOMIC DNA FROM hESCs USING PHENOL/CHLOROFORM EXTRACTION This protocol describes an alternative method of isolating genomic DNA from hESCs grown in multi-well plates. A phenol/chloroform extraction is included to improve the quality of the genomic DNA. This approach should be considered if no hESC clones containing the desired genetic modification are identified when DNA is isolated using the techniques described in Support Protocol 2, or it is already known that the genomic fragment is difficult to amplify in the PCR screening strategy. This protocol can also be used to give high-quality DNA directly from cells on the primary plate if all the wells containing hESCs are confluent, and if there are fewer than 48 clones to screen, saving ∼1 week of culturing time (as described in Support Protocol 1). If there are more than 48 clones, it is difficult for one person to process all of the samples using this procedure.

Materials 48-well tissue culture plates containing confluent hESC colonies (Support Protocol 1, step 13) or microcentrifuge tubes containing clumps of hESCs (Support Protocol 1, step 13, annotation) DNA lysis buffer containing 200 μg/ml proteinase K (see recipe) Phenol/chloroform/isoamyl alcohol (25:24:1) solution saturated with 10 mM Tris·Cl, pH 8.0/1 mM EDTA 70 %(v/v) and 100% (v/v) ethanol Trypsin/EDTA buffer (TE buffer; see recipe) Generation of hESC Reporter Knock-In Lines


1.5-ml microcentrifuge tubes Vortex mixer Microcentrifuge 55◦ C incubator Current Protocols in Stem Cell Biology

1. Aspirate the medium from the hESC clones growing on the 48-well plates labeled “DNA.” If the hESCs are in microcentrifuge tubes (from Support Protocol 1), pellet the cells by centrifuging 3 min at 480 × g, 4◦ C, before removing the medium.

2. Add 100 μl DNA lysis buffer containing 200 μg/ml proteinase K to each clone and incubate for ∼3 hr at 55◦ C. Alternatively, incubate overnight at 37◦ C.

3. Transfer the contents of each well to individual 1.5-ml microcentrifuge tubes. Label the tubes such that the corresponding hESC clone can be identified on the “Maintenance” multi-well plate. If the hESCs are already in microcentrifuge tubes, skip step 3 and proceed to step 4.

4. Add an equal volume of phenol/chloroform/isoamyl alcohol solution to each tube, vortex to mix well, and then centrifuge the tubes 10 min at 10,000 × g, room temperature. 5. Carefully transfer the top layer containing the DNA to a new 1.5-ml microcentrifuge tube containing 250 μl of 100% ethanol. Mix the solutions until the DNA precipitates. 6. Pellet the DNA by microcentrifuging 10 min at maximum speed, room temperature. 7. Carefully remove the supernatant and wash the DNA pellet with 250 μl of 70% ethanol. Ensure all ethanol is removed by leaving the DNA pellet to air dry for 10 min. 8. Resuspend each sample in 50 μl TE buffer, and incubate for 3 to 4 hr at 55◦ C to allow the DNA to dissolve. 9. Store the DNA samples for at least 6 months at 4◦ C.

PCR IDENTIFICATION OF TARGETED hESC CLONES To identify targeted hESC clones, a PCR screening strategy is employed. This technique enables a large number of clones to be screened rapidly. The PCR screen is designed to amplify a novel junction fragment created by the correct homologous recombination event. One primer should anneal to sequences located in either the positive selection cassette or in the reporter gene (see Table 5B.1.1 for a list of recommended primers), while the second primer should prime from the target chromosomal sequences just beyond the homologous sequences used in the targeting vector (Fig. 5B.1.5). This method is also utilized to identify hESC clones in which the positive selection cassette has been excised from the genome (Basic Protocol 2), and to identify targeted lines following clonal isolation (Support Protocol 4). The robustness of the PCR amplification is in part related to the distance between the two primers and the composition of the DNA sequence being amplified. It is recommended that the amplified product be between 2 and 4 kb in length, and not contain long GC stretches. The addition of DMSO to the reaction may also assist in the amplification. Sequences of a similar size can be amplified from the wild-type locus (for example using primer ’b’ in Fig. 5B.1.5 and another primer binding to the wild-type locus ∼4 kb upstream) to assist in the optimization of the PCR reaction. The authors recommend using Platinum Taq DNA Polymerase High Fidelity (Invitrogen), and have found that this enzyme mixture regularly amplifies 1- to 5-kb sequences from genomic DNA using the reaction mixture listed in Table 5B.1.2.

SUPPORT PROTOCOL 3



Supplement 11

Table 5B.1.1 Primers Known to Anneal to Fluorescent Marker Genes and DrugResistance Cassettes

Gene

Sequences are 5 to 3

Annealing temperature

Direction of amplification

GFP

GTGCTGCTGCCCGACAACCACTAC CCGGTGAACAGCTCCTCGCCCTTGC

60◦ C 62◦ C

FWD (5 to 3 ) REV (3 to 5 )

RFP

CACAACACCGTGAAGCTGAAGGTGAC GTCACCTTCAGCTTCACGGTGTTGTG

65◦ C 65◦ C

FWD (5 to 3 ) REV (3 to 5 )

neo

CGATGCCTGCTTGCCGAATATCATG

60◦ C

FWD (5 to 3 )

hph

CTCCGCATTGGTCTTGACCAACTC

60◦ C

FWD (5 to 3 )

targeting vector

reporter

pos. select

exon 1

wild-type allele

exon 2

a targeted allele

reporter

b

pos. select

exon 2

Cre Recombinase c reporter

b exon 2

Figure 5B.1.5 A schematic representation of homologous recombination between the targeting vector and the wild-type allele. The wild-type allele represents a typical target gene containing two exons. The targeting vector contains sequences homologous to the genomic locus (orange lines and rectangles), as well as sequences encoding a reporter gene (green arrow), loxP sites (red triangles), and a positive selection cassette (blue rectangle). Homologous recombination between the wild-type allele and the targeting vector replaces a segment of exon 1 in the wild-type allele. A PCR assay is used to detect gene-targeted clones with primers (black arrows) corresponding to sequences within the positive selection cassette (a) and the endogenous wild-type allele (b). Only correctly targeted alleles will yield a PCR product. Following expression of Cre recombinase in the cells, the positive selection cassette is removed from the targeted allele. The loss of the positive selection cassette is confirmed by a second PCR using the same endogenous primer (b) and a primer corresponding to sequences within the reporter gene (c).

If any correctly targeted hESC clones are identified from the PCR screen, these cell lines should be maintained as colonies that are mechanically passaged, and stocks of each line frozen in liquid nitrogen. Genetically manipulated hESCs that are kept and grown as colonies retain their stem cell characteristics and are indistinguishable in appearance from the parental lines (Costa et al., 2005; Davis et al., 2008a).

Materials


PCR master mix (see Table 5B.1.2) containing: Autoclaved, distilled water 10× High Fidelity PCR Buffer (Invitrogen) 10 mM dNTP mixture (Sigma) 50 mM MgSO4



Forward & Reverse Primers (see Table 5B.1.1) DMSO Platinum Taq High Fidelity DNA polymerase (Invitrogen) Genomic DNA from the hESC clones (Support Protocol 2 or the Alternate Protocol) hESC medium (see recipe) Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) 0.2-ml nuclease-free PCR tubes 1.5-ml nuclease-free microcentrifuge tubes Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Microcentrifuge DNA thermal cycler Gelatinized organ culture dishes Additional reagents and equipment for analyzing the amplification products by agarose gel electrophoresis (Voytas, 2001) and maintaining and expanding hESCs by both mechanical and enzymatic passaging (UNIT 1C.1) Set up and run the PCR 1. Organize the PCR tubes in the same layout as the DNA samples, on ice. 2. Determine the total number of DNA samples to screen. Include in this total both negative and positive control samples, if available. A possible negative control DNA sample is genomic DNA from nontransfected hESCs. Generally, a positive control DNA sample is not included. However, if a hESC line has previously been successfully targeted at that genomic locus, this can serve as a positive control.

3. Make a PCR master mix in a 1.5-ml nuclease-free microcentrifuge tube, on ice. A suggested cocktail is listed in Table 5B.1.2. Optimal concentrations for the PCR master mix will depend on the primer pair used and the DNA sequence being amplified, and must be determined empirically. This reagent assembly is best performed in a dedicated PCR preparation area using pipettors that have not been used to aliquot template DNA.

4. Aliquot 18 μl of the master mix into each PCR tube. Table 5B.1.2 PCR Master Mix

Amounta

Component

(n + 1) × 12.9 μl

dH2 O 10× High Fidelity PCR buffer 50 mM MgSO4

(n + 1) × 1.2 μl (n + 1) × 1 μl

DMSO 10 mM dNTPs 250 ng μl

(n + 1) × 2 μl

−1

forward primer

(n + 1) × 0.4 μl (n + 1) × 0.2 μl

250 ng μl−1 reverse primer

(n + 1) × 0.2 μl

5 U μl−1 Platinum Taq High Fidelity DNA polymerase

(n + 1) × 0.1 μl

Total

(n + 1) × 18 μl

a n equals the total number of reactions to be performed, including the positive and negative control

samples.



Supplement 11

5. To each PCR tube, add 2 μl of the appropriate genomic DNA sample. The DNA solutions are usually between 50 and 200 ng μl−1 and, in the 48-well plates, are often very viscous. These samples need to be mixed well with the pipet tip before removing 2 μl. To assist in tracking the PCR tubes into which the DNA samples have been added, leave the pipet tips containing the template DNA in the tube until all DNA samples have been aliquoted. Use a different pipettor to aliquot the DNA samples from the one used to aliquot the PCR master mix reagents.

6. Seal and briefly microcentrifuge the tubes 30 sec at 100 × g, room temperature. 7. Place tubes inside a thermal cycler, and enter cycle conditions. Commonly used amplification conditions are: 1 cycle:

3 min

30 to 40 cycles: 20 sec 30 sec 1 min (per kb of sequence being amplified) 1 cycle: 10 min

94◦ C 94◦ C 50◦ to 62◦ C 68◦ C

(initial denaturation) (denaturation) (annealing) (extension)

68◦ C

(final extension).

8. Visualize the PCR products by agarose gel electrophoresis (Voytas, 2001), and identify the hESC clones that have the desired genetic modification. If none of the hESC clones analyzed appear to be targeted, a second PCR should be performed using primers to amplify a similar-sized fragment in the same genomic region from the wild-type allele. A correct sized product obtained using this second set of primers indicates that it is unlikely that the absence of a PCR product from the screening PCR was a consequence of poor DNA template quality or low DNA concentration. If the primers specific to the wild-type allele fail to amplify the DNA fragment, further purification of the hESC DNA samples and/or optimization of the PCR protocol will be required prior to repeating the screening PCR.

Transfer the correctly targeted hESC clones onto organ culture dishes 9. Leave the hESC clones with the correct genetic modification to expand on the “Maintenance” 48-well plates, until the wells are ∼80% confluent (Fig. 5B.1.4B) or the colonies are beginning to differentiate. Change the hESC medium on the wells daily. Usually the hESC clones will be at this stage of growth by the time the PCR screening is complete.

10. One day before transferring the hESC clones, plate mitotically inactivated MEFs onto the center well of gelatinized organ culture dishes at a density of 6 × 104 per cm2 in 1 ml of MEF medium. Prepare enough organ culture dishes with MEFs, such that pieces from each targeted hESC clone can be plated onto two organ culture dishes.

11. On the day of transfer, replace the MEF medium on the organ culture dishes with 1 ml of hESC medium. 12. Mechanically fragment a hESC clone to be moved using the same technique described in step 12 of Support Protocol 1, and distribute the pieces of the clone across two organ culture dishes.


13. Repeat the above step with all genetically modified hESC clones that are to be transferred to organ culture dishes. Repeat the PCR screen on all positive clones that are identified to verify that the correct clone has been transferred from the maintenance plate.



14. Maintain the reporter gene knock-in hESC lines as colonies that are mechanically passaged (UNIT 1C.1). Also expand the hESCs as a large-scale expansion by enzymatic passaging (UNIT 1C.1) for deletion of the positive selection cassette from the targeted lines (Basic Protocol 2). All targeted knock-in hESC lines generated at this stage should have stocks frozen in liquid nitrogen. A suggested method of cryopreservation is described elsewhere (Reubinoff et al., 2001).

REMOVING THE POSITIVE SELECTION CASSETTE FROM GENETICALLY MODIFIED hESCs BY TRANSIENT EXPRESSION OF Cre RECOMBINASE

BASIC PROTOCOL 2

Targeting vectors contain positive selection cassettes to facilitate the isolation of stably transfected cells. Following the isolation and identification of targeted lines, selection cassettes are removed because their continued presence may cause a number of undesirable effects. In genetically modified mice, the retention of the positive selection cassette can interfere with the expression of neighboring endogenous genes (Pham et al., 1996; Scacheri et al., 2001), while in reporter knock-in hESC lines it can result in the misexpression of the reporter (either by silencing or activating expression of the reporter; R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.). The removal of the positive selection cassette also offers the opportunity to subsequently retarget the remaining wild-type allele and to generate a homozygous knockout hESC line using the original targeting vector and drug selection strategy. If the positive selection cassette is flanked by loxP sites, expression of Cre recombinase in the genetically modified cells catalyzes the excision of the DNA sequence between the loxP sites (Sauer, 1993). This protocol describes a method for the transient transfection of a circular pEFBOS-creIRESpuro vector into hESCs using the lipofection reagent FuGENE 6 (Roche), and is an alternative to the transduction of a recombinant-modified Cre recombinase protein into the cells (Nolden et al., 2006). The transfected cells constitutively express both Cre recombinase and puromycin N-acetyltransferase and are selected for by the addition of the antibiotic puromycin to the hESC medium for 48 hr. This short selection period enriches for cells that have been transiently transfected with the pEFBOS-creIRESpuro vector and also allows sufficient time for Cre-mediated deletion of the antibiotic-resistance cassette to occur. Approximately 12 days after selection, the hESC colonies that have formed can be picked and expanded as described in Support Protocol 1. DNA is then extracted (Support Protocol 2) and the loss of the positive selection cassette is confirmed by PCR (Support Protocol 3).

Materials Gene-targeted hESCs containing a loxP-flanked positive selection cassette in gelatinized 75-cm2 tissue culture flasks between enzymatic passages 5 and 10 co-cultured with MEFs pre-seeded at a density of 1.5 × 106 /75-cm2 flask Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) hESC medium (see recipe) 60-mm gelatinized tissue culture dishes (see recipe) seeded with 3 × 104 /cm2 mitotically inactivated MEFs Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) FuGENE 6 Transfection Reagent (Roche) DMEM/F12 (Invitrogen) pEFBOS-CreIRESpuro expression vector (GenBank accession number EU693012; available on request from the authors’ laboratory; e-mail request to [email protected] or [email protected])



Supplement 11

Puromycin solution (Sigma), 10 mg/ml Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips 37◦ C incubator 15- and 50-ml sterile centrifuge tubes Refrigerated centrifuge 1.5-ml microcentrifuge tubes Additional reagents and equipment for performing a cell count (Phelan, 2006; UNIT 1C.3), transferring colonies to 48-well tissue culture plates (Support Protocol 1), extracting DNA from colonies (Support Protocol 2), and screening extracted DNA by PCR (Support Protocol 3) Seed feeders onto 60-mm dishes (day −2) 1. Supplement five 60-mm dishes with 2 ml gelatin to ensure that the gelatin coats the surface. 2. Allow to stand for 30 min at room temperature. 3. Aspirate gelatin and seed 6 × 105 feeders/dish in 4 ml MEF medium. 4. Store in a humidified incubator at 37◦ C, 5% CO2 until required.

Enzymatically passage genetically modified hESCs (day −1) 5. Harvest the genetically modified hESCs cultured in a 75-cm2 flask. Aspirate the hESC medium and rinse the flask with 5 ml CMF-PBS. Add 2 ml of trypsin or TrypLE Select to the flask and ensure that the dissociation solution coats the surface of the cells. Place the flask 4 hr at 37◦ C for 4 min and dislodge the hESCs from the flask with gentle tapping. 6. Add 8 ml of hESC medium to the flask and transfer the resuspended hESCs to a 15-ml centrifuge tube. 7. Pellet the cells by centrifuging the tube 3 min at 480 × g, 4◦ C, and remove the supernatant. 8. Resuspend the hESC pellet in 5 ml of fresh hESC medium and perform a cell count (Phelan, 2006; UNIT 1C.3). Subtract the number of MEFs (∼0.75 × 106 ) from the count to determine the total number of hESCs. Generally, a semi-confluent 75-cm2 flask will contain ∼4 × 106 hESCs, which is enough cells to seed five 60 mm-dishes with ∼0.8 × 106 hESCs per dish.

9. Transfer 4 × 106 hESCs to a 50-ml centrifuge tube. Add additional hESC medium so that the cells are resuspended in a total volume of 20 ml. 10. Distribute 4 ml of the hESC suspension into each of the five 60-mm dishes seeded the day before (steps 1 to 4) with MEFs at a density of 3 × 104 cells/cm2 . 11. Return the dishes to a humidified incubator at 37◦ C, 5% CO2 , and leave the hESCs to attach overnight.

Transfect the Cre recombinase expression vector into the hESCs (day 0) 12. Approximately 2 hr before performing the transfection, aspirate the medium on the 60-mm dishes and supplement the cells with 4 ml fresh hESC medium.


The dishes should be no more than 70% confluent with hESCs. If the cells are overconfluent, the hESCs may not transfect optimally with FuGENE. In addition, we have observed a reduction in efficacy of puromycin in eliminating untransfected hESCs if the hESCs are overconfluent at the time of antibiotic selection.



13. Add 30 μl of FuGENE 6 Transfection Reagent directly to 470 μl DMEM/F12 in a sterile 1.5-ml microcentrifuge tube. Mix by flicking the microcentrifuge tube and incubate the complex for 5 min at room temperature The manufacturer advises that the undiluted FuGENE 6 Transfection Reagent should not come into contact with the walls of the microcentrifuge tube, as this can adversely affect the transfection efficiency.

14. Transfer 100 μl of the FuGENE 6/DMEM/F12 mixture into another microcentrifuge tube labeled “negative control.” This solution is applied to the fifth 60-mm dish that serves as a negative control.

15. Pipet 4 μl of a 1 μg/μl stock solution of pEFBOS-creIRESpuro plasmid DNA into the 400 μl FuGENE 6/DMEM/F12 mixture. Tap the microcentrifuge tube to mix the contents and leave at room temperature for a further 40 min. Incubate the 100 μl negative control centrifuge tube for the same period of time. The pEFBOS-creIRESpuro plasmid preparation must be pure and endotoxin-free. Transfection-grade plasmid preparation can be isolated using the Qiagen plasmid purification kits.

16. Into four of the hESC-containing 60-mm dishes, add 100 μl of the FuGENE/DNA complex mixture dropwise. Add the negative control into the fifth 60-mm dish. Swirl the dishes to ensure distribution over the entire surface, label all dishes appropriately, and return to the incubator. A negative control dish of hESCs for the FuGENE transfection is a good indicator of the kinetics and completeness of cell death in response to puromycin.

Select transiently transfected hESCs (day 1) 17. Supplement 50 ml of hESC medium with puromycin to a final concentration of 2 μg/ml. The optimal concentration of puromycin for selection may vary for different hESC lines. A titration should be performed to determine the minimum concentration required to eliminate untransfected hESCs within 48 hr of addition. For 2 days of selection, 50 ml of hESC medium containing puromycin should be sufficient stock. Discard any unused stock after 2 days.

18. Apply 5 ml of the selection medium to each of the five 60-mm dishes between 24 to 36 hr after FuGENE transfection. The hESCs should be ∼90% confluent.

19. Maintain puromycin selection for 48 hr and change the medium daily. Frequent medium changes are necessary during selection to remove the dead cells. After 48 hr, all the hESCs in the control dish should have died. If the dish still contains viable hESCs, either the concentration of puromycin was not sufficient to eliminate untransfected hESCs, or the dishes were too confluent with hESCs when selection was started.

20. Following puromycin selection, return to culturing the cells in hESC medium that does not contain puromycin. If the MEFs preseeded on the 60-mm dishes are puromycin-sensitive, they will need to be replaced once puromycin selection is stopped. Supplement each dish with ∼0.6 × 106 MEFs resuspended in hESC medium. If the MEFs are puromycin-resistant, only supplement with additional MEFs when gaps in the MEF layer appears. Try to maintain a MEF density of ∼2 × 104 viable MEFs/cm2 on the dishes.

21. Allow the colonies to grow for ∼12 days, changing the hESC medium daily.



Supplement 11

22. Once the colonies are ∼2-mm in diameter, transfer a minimum of 24 undifferentiated colonies to a 48-well tissue culture plate as described in Support Protocol 1 (steps 2 to 8). 23. Then extract DNA from these colonies (Support Protocol 2) and screen it by PCR (Support Protocol 3) to confirm that the positive selection cassette has been removed from the hESCs. This is achieved by choosing a 5 primer that anneals to sequences within the reporter gene, and a 3 primer to sequences 3 of the antibiotic-selection cassette (Fig. 5B.1.5). A genomic DNA sample from targeted hESCs that still contain the selection cassette can be included in the screen to visualize the difference in size between PCR products still containing the positive selection cassette and those in which the cassette has been excised. Additional PCR screens using primers specific to the Cre recombinase expression vector to verify that the plasmid did not integrate into the genome of the resulting colonies, and a PCR using primers specific to the antibiotic-resistance cassette to exclude the presence of residual unexcised cells, should be performed. The sensitivity of the hESCs to geneticin and puromycin can also be confirmed later by re-exposing an aliquot of the cells to the selection agents. Two or three of these reporter knock-in hESC clones that have had the positive selection cassette removed should be returned to organ culture dishes for maintenance and expansion. These lines should also be cloned as described in Support Protocol 4 to ensure that the targeted hESC lines used in future applications contain a targeted locus in which the antibiotic selection cassette has been excised. SUPPORT PROTOCOL 4

CLONAL ISOLATION OF hESCs BY SINGLE-CELL DEPOSITION FLOW CYTOMETRY The procedures for generating a targeted hESC line and removing the positive selection cassette support the clonal growth of hESCs. However, they do not necessarily exclude the possibility that the resulting hESC colonies arose from more than one cell. This protocol uses flow cytometry to derive single-cell clones from the existing parental hESC lines. Despite the low cloning efficiency of hESCs reported in the literature (Sidhu and Tuch, 2006), we routinely achieve a single-cell cloning frequency of 4% to 8% using hESCs adapted to enzymatic passage as we have described in UNIT 1C.1. Viable hESCs are selected by size gating and exclusion of the dye, propidium iodide (PI), and single cells are deposited directly into individual wells of 96-well plates using a flow cytometer. Correct gene targeting and absence of the positive selection cassette is confirmed in the resulting clones by PCR. It is possible to merge this protocol with the procedure to remove the positive selection cassette (Basic Protocol 2), reducing the time required to generate the final targeted hESC line. A description of this integrated procedure is provided elsewhere (Davis et al., 2008b). Subcloning ensures that the resulting targeted hESC lines consist of cell populations with identical genetic constitutions. This method is also a useful approach for ensuring a homogeneous diploid cell population, and selecting hESC lines with a uniform, undifferentiated morphology.

Materials


0.1% (w/v) gelatin solution (see recipe) Mitotically inactivated, irradiation-treated (UNIT 1C.3) mouse embryonic fibroblasts (MEFs; Conner, 2000) MEF medium (see recipe) Genetically modified hESCs generated from Basic Protocol 2 in 75-cm2 tissue culture flasks between enzymatic passages 5 and 10



75-cm2 gelatinized tissue culture flasks (see recipe) seeded with mitotically inactivated MEFs at 1 × 104 /cm2 for passaging hESCs prior to cloning hESC medium (see recipe) Recombinant human FGF2 (see recipe) Phosphate-buffered saline without CaCl2 and MgCl2 (CMF-PBS; Invitrogen) Trypsin (see recipe) or TrypLE Select cell dissociation enzyme (Invitrogen) Propidium iodide (PI) solution (see recipe) Flat-bottomed 48-well tissue culture plates, gelatinized and seeded with mitotically inactivated MEFs at 0.75 × 106 /plate Liquid nitrogen 96-well flat-bottom tissue culture-treated plates and lids 37◦ C, 5% CO2 incubator 15-ml sterile centrifuge tubes Refrigerated centrifuge 5-ml sterile round-bottom polystyrene FACS tubes (12 × 75–mm) with 35-μm cell-strainer caps and with snap lids (Falcon) Parafilm M (Pechiney Plastic Packaging) or equivalent Flow cytometer with single-cell deposition function, e.g., FACSVantageSE-DiVa system (Becton Dickinson) or equivalent Inverted microscope Gilson pipettors (John Morris Scientific) or equivalent, with sterile (plugged) tips Stereomicroscope Additional reagents and equipment for propagating hESCs in bulk culture (UNIT 1D.3), extracting DNA from colonies (Support Protocol 2), screening extracted DNA by PCR (Support Protocol 3), maintaining and expanding the genetically modified lines (UNIT 1C.1), and for cryopreserving hESCs (Reubinoff et al., 2001) Feeder reduce the hESCs to be subcloned and prepare the 96-well plates (day −1) 1. Add ∼50 μl of 0.1% gelatin solution to each well on ten 96-well plates. Leave the plates at room temperature for 15 min. 2. Aspirate the gelatin solution from the wells. 3. Resuspend MEFs in MEF medium such that the final concentration is 2 × 105 MEFs/ml. Aliquot 50 μl of the MEF-containing medium into each well. The final density of MEFs is ∼1 × 104 MEFs/well.

4. Passage hESCs for subcloning into a 75-cm2 flask containing MEFs seeded at a density of 1 × 104 /cm2 (UNITS 1C.1 & 1D.3). Harvest the hESCs as described in steps 1 to 3 of Basic Protocol 2. Resuspend the pelleted hESCs in hESC medium and transfer three-quarters of the cell suspension to the 75-cm2 flask. To maintain the hESC line, the remaining cells can be plated into another tissue culture flask containing MEFs at the normal density (2 × 104 MEFs/cm2 ). The passaging of the hESCs onto low-density MEFs increases the hESC:MEF ratio, improving the number of hESC subclones obtained following single-cell deposition.

5. Incubate both the 96-well plates and the flasks containing the passaged hESCs in a humidified incubator at 37◦ C, 5% CO2 overnight.

Subcloning of the gene-targeted hESC line (day 0) 6. Supplement 300 ml of hESC medium with additional FGF2 so that the final concentration of FGF2 is 40 ng/ml.



Supplement 11

The higher concentration of FGF2 supports the growth and expansion of the hESCs, and also suppresses the spontaneous differentiation that can occur during the early establishment of the single-cell clones.

7. Aspirate the MEF medium from the wells of the multi-well plates, and aliquot 100 μl of hESC medium containing 40 ng/ml FGF2 into each well. Return the plates to the incubator until required for use with the flow cytometer.

Harvest hESCs 8. Harvest the hESCs to be subcloned. Briefly, remove the medium from the 75-cm2 flask, rinse the cells with 5 ml CMF-PBS, add 3 ml of the dissociating agent (either TrypLE Select or trypsin) and return the flask to the 37◦ C incubator for 4 to 5 min. 9. Gently tap the base of the flask to dislodge the hESCs and add 10 ml hESC medium (without the additional FGF2) to the flask. Mix and transfer this cell suspension to a 15-ml centrifuge tube. 10. Pellet the cells by centrifuging the tube for 3 min at 480 × g, 4◦ C. 11. Remove the supernatant and resuspend the hESCs in 1 ml of hESC medium (without the additional FGF2). 12. Filter the hESCs by passing the cell suspension through a 35-μm cell-strainer cap attached to a sterile FACS tube. To help the dissociated hESCs pass through the cell strainer, cover the caps with a small square of Parafilm and centrifuge the tube for 3 min at 480 × g, 4◦ C, to pellet the cells. This step removes cell clumps and cellular debris and ensures that the hESCs form a single-cell suspension. Covering the caps the Parafilm helps to keep the cells sterile.

13. Return the tube to the tissue culture hood and discard the Parafilm and cell strainer cap. 14. Carefully aspirate the supernatant and resuspend the hESCs in 1.5 ml of hESC medium supplemented with 40 ng/ml FGF2. To this mixture, also add 15 μl of 100 μg/ml propidium iodide (PI) solution. Flick the tube to mix. 15. Recap the FACS tube with a sterile FACS tube cap and store at 4◦ C or on ice until required. Sealing the FACS tube prevents desiccation and keeps the cell suspension sterile.

16. Identify the viable hESCs by size gating and exclusion of PI on a flow cytometer whose lines have been sterilized. Deposit single cells directly into each well of the ten 96-well plates. Although not routinely used in our laboratory, the cloning efficiency of the hESCs can be further improved by treating the cells with the Rho-associated kinase inhibitor (ROCKi), Y-27632 (Watanabe et al., 2007). We have had success by adding the inhibitor at 2 μM to the hESC medium in the 96-well plates at the time of sorting.

17. Return the plates to a humidified incubator at 37◦ C, 5% CO2 . 18. Four days after seeding the plates with hESCs, supplement the medium in each well with an additional 50 μl of hESC medium containing 40 ng/ml FGF2. We do not advocate the continued inclusion of ROCKi in the culture medium.


Identify wells containing hESC colonies (day 8) 19. Use an inverted microscope to identify wells containing viable hESC colonies. Carefully aspirate the medium from these wells and replace with 100 μl of hESC



medium containing 40 ng/ml FGF2, as well as fresh MEFs at a concentration of 3 × 104 MEFs/ml. 20. Allow the colonies to continue growing. Replace the hESC medium on the wells containing viable hESC colonies daily. When the colonies are ∼2-mm in diameter, the hESCs can be transferred to and expanded on 48-well tissue culture plates as described in step 21. However, if the colonies begin to differentiate before they reach that size, they should be transferred sooner.

Transfer the hESC colonies onto 48-well plates (between days 16 to 18) 21. Determine the total number of viable hESC colonies on the 96-well plates. On the day before moving the colonies, seed enough 48-well plates with ∼0.75 × 106 MEFs/gelatinized plate (∼2 × 104 /cm2 ) so that each hESC colony can be transferred to an individual well. Incubate the 48-well plates overnight at 37◦ C, 5% CO2 to allow the MEFs to attach. Generally, we observe between 40 to 80 colonies spread over the ten 96-well plates.

22. Replace the medium on the 48-well plates with 200 μl of hESC medium supplemented with 10 ng/ml FGF2 per well. The concentration of FGF2 in the hESC medium can be returned to 10 ng/ml from this stage onwards.

23. Fragment a hESC colony into multiple small cell clumps with a 200-μl plugged tip attached to a pipettor. Transfer these pieces into a well on one of the 48-well plates. This can be performed macroscopically and the well on the 96-well plate checked afterwards under a stereomicroscope to confirm that the entire hESC colony detached and was transferred.

24. Discard the pipet tip after harvesting each colony. 25. Repeat steps 23 and 24 to transfer the remaining hESC colonies growing on the 96-well plates. The 96-well plates can be discarded once all the viable hESC colonies have been transferred.

Maintain and expand the clones 26. Maintain replicate and expand the hESC colonies as described in steps 8 to 17 in Support Protocol 1. 27. Extract DNA from these colonies (Support Protocol 2) and perform a PCR screen (Support Protocol 3) to confirm the subclones are correctly targeted, have the positive selection cassette excised from the genome, and did not integrate Cre recombinase into the genome (Basic Protocol 2, step 21, annotation). Discard any subclones that are negative for this screen. 28. From the remaining subclones, choose two or three that are growing well and appear undifferentiated in culture, and return these colonies to organ culture dishes. 29. Maintain and expand these genetically modified lines using the protocols described in UNIT 1C.1. 30. Confirm that these subclones have maintained a stem cell phenotype and a normal karyotype. 31. Cryopreserve stocks of these lines (Reubinoff et al., 2001). Generic Manipulation of Stem Cells


Supplement 11


DNA lysis buffer 100 mM Tris·Cl, pH 8.0 200 mM NaCl 5 mM EDTA, pH 8.0 Add 0.2% (w/v) SDS powder (Sigma) last At time of use, dispense the required volume and add 200 μg/ml proteinase K (see recipe) to the solution Store the solution without proteinase K at room temperature indefinitely Gelatin, 0.1% (w/v) Add 0.5 g of gelatin powder (from porcine skin; Sigma) to 500 ml distilled water and autoclave to dissolve and sterilize. Store up to 6 months at room temperature.

Gelatinization of plates and flasks Prior to addition of MEFs, add enough 0.1% (w/v) gelatin solution (see recipe) to cover the base of all plates/flasks. Let stand for 10 min at 37◦ C to coat the surface and remove by aspiration immediately prior to addition of MEFs.

hESC medium DMEM/F12 (Invitrogen) containing: 20% (v/v) Knockout Serum Replacement (Invitrogen) 10 mM non-essential amino acids (Invitrogen) 2 mM L-glutamine or GlutaMaxI (Invitrogen) 1× penicillin/streptomycin (200× stock; Invitrogen) 100 μM 2-mercaptoethanol 10 ng/ml FGF2 (see recipe) Filter sterilize using a 0.22-μm Stericup filtration unit (Millipore) Store up to 1 week at 4◦ C MEF medium DMEM (4.5 g/liter glucose, without L-glutamine and sodium pyruvate; Invitrogen) containing: 10% (v/v) heat-inactivated fetal bovine serum (FBS) 2 mM L-glutamine (Invitrogen) 1× penicillin/streptomycin (200× stock; Invitrogen) Filter sterilize and store up to 4 weeks at 4◦ C Propidium iodide (PI) Dissolve PI (Sigma) in CMF-PBS to a final stock concentration of 100 μg/ml (100× stock). Filter sterilize and store for at least 12 months at 4◦ C.

Proteinase K Dissolve proteinase K powder (Sigma) in H2 O to a final stock concentration of 20 μg/μl (20 mg/ml; 100×). Dispense into 1-ml aliquots and store for at least 12 months at –20◦ C. Generation of hESC Reporter Knock-In Lines

Recombinant human basic fibroblast growth factor (FGF2) Reconstitute lyophilized rhFGF2 (PeproTech) to a final stock concentration of 10 μg/ml in CMF-PBS. Store up to 6 months at –80◦ C.



Soybean trypsin inhibitor Dissolve the soybean trypsin inhibitor stock powder (Sigma) in CMF-PBS to a final concentration of 1 mg/ml. Filter sterilize and dispense into 5- or 10-ml aliquots and store up to 12 months at −20◦ C. Once thawed, the solution can be stored for 1 to 2 weeks at 4◦ C.

Tris/EDTA (TE) buffer, pH 8.0 10 mM Tris·Cl, pH 8.0 1 mM EDTA, pH 8.0 Store the solution at room temperature indefinitely Trypsin, 0.125% (w/v) Supplement Trypsin/EDTA [0.25% (w/v) trypsin EDTA. 4Na, Invitrogen] with 2% (v/v) chicken serum (Hunter antisera). Decant into 5-ml aliquots and store up to 12 months at −20◦ C. To use, thaw an aliquot and add an equal volume of CMF-PBS. Once thawed, the solution can be stored up to 4 weeks at 4◦ C.

COMMENTARY Background information Human embryonic stem cells are derived from the inner cell mass of the pre-implantation blastocyst stage embryo (Thomson et al., 1998; Reubinoff et al., 2000). These cells exhibit two key characteristics. Firstly, they can be maintained and expanded in vitro for extended periods of time as undifferentiated cells while preserving their original karyotype. Secondly, they are pluripotent, and therefore have the capacity to differentiate into various cell types representing the three germ layers, both in vivo and in vitro (Amit et al., 2000). The ability to transform hESCs into multiple lineages in culture provides opportunities to examine human embryonic development in vitro, generate specific cells and tissues for therapies or drug screening, and identify chemical compounds that influence a specific developmental process. The identification of a cohort of lineagespecific markers assists in the directed differentiation of hESCs towards specific cell types. When appropriate antibodies for specific cellsurface markers are available, fluorescenceactivated cell sorting (FACS) may be used to purify viable hESC derivatives. Where antibodies for suitable cell surface markers are unavailable, or the lineage-specific markers are intracellular, reporter genes can be targeted to loci whose expression marks critical developmental milestones, facilitating the isolation of viable cell populations that would otherwise be inaccessible. These purified subpopulations can then be further differentiated or expanded in vitro, or transplanted and tracked

in vivo. These approaches have contributed significantly to our understanding of lineage specification in differentiating mouse ESCs (Fehling et al., 2003; Ying et al., 2003; Ng et al., 2005; Micallef et al., 2005, 2007; Gadue et al., 2006). Until recently, similar strategies have been unachievable with hESCs, partly due to suboptimal culturing conditions leading to poor transfection and single-cell cloning efficiencies. However, several studies have now described the introduction of selectable markers into hESCs using a variety of different approaches (Giudice and Trounson, 2008, and references therein). These include using vectors or viruses to randomly integrate reporter genes regulated by lineage-specific promoter fragments into hESCs. While these methods can introduce new genetic material into the genome at relatively high frequencies, there are several caveats associated with random integration, the foremost being that expression of the reporter may not faithfully reflect expression of the endogenous gene. Other concerns include the possibility of disrupting normal gene functions, and that elements other than the promoter fragments included in the construct may be required for accurate expression of the reporter. Furthermore, unlike mouse ESC lines, in which transgenic lines can be validated by examining reporter gene expression patterns in chimeric mice, validation of analogous hESC lines relies on examination of reporter expression in vitro or the more limited in vivo setting afforded by xenogenic transplants.



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Gene targeting utilizes the cellular DNA repair machinery to integrate transgenes, such as reporter genes, by homologous recombination into a specific site in the genome. The reporter gene is regulated in the same manner as the endogenous gene from the remaining wild-type allele and is thus expressed at the same time and in the same cells. Therefore, although homologous recombination is technically challenging, it produces expression patterns that more accurately reflect native gene expression and is preferable to random integration for the production of reporter cell lines. While electroporation is the chosen method for gene targeting in mouse ESCs, initially it was considered that this technique was unlikely to be successful in hESCs due to the resulting high mortality (Eiges et al., 2001). Modifications to both the culturing of hESCs and to the standard mouse ESC electroporation protocol have formed the basis of a successful targeting method (Costa et al., 2007). While it is possible to obtain homologous recombinants in hESCs using cationic reagents, this occurs at a very low frequency (10−8 ; Urbach et al., 2004). Using the electroporation procedure described in this unit, the stable transfection frequency is consistently between 2 × 10−6 and 5 × 10−5 (i.e., 20 to 500 colonies per 107 transfected cells), of which 1.3% to 14.6% of the stable transfectants are correctly targeted, depending on the locus and the specific hESC line (Costa et al., 2007). While the stable transfection frequency in hESCs is around 100-fold lower than what is usually observed in mESCs (Vasquez et al., 2001), the frequency of homologous recombination within stable transfectants appears comparable between the two species. During the differentiation of hESCs, a panel of markers are generally used to screen and identify lineage-specific cell types. Engineering dual or multiple reporter knock-in hESC lines would enable the identification of pools of cells that share common markers and improve our understanding of the molecular and cellular mechanisms that govern lineage specification. Results from these lines of analyses can subsequently be utilized to differentiate unmodified hESC lines to cell types with potential clinical applications.

Critical Parameters and Troubleshooting Generation of hESC Reporter Knock-In Lines

Gene targeting strategy The generation of reporter knock-in hESC lines is a challenging task, as illustrated by the

limited number of reports describing successful gene targeting in hESCs. In our laboratory, we aim to generate several independent genetargeted clones in more than one hESC line in order to demonstrate the generalizability of the experimental findings. The planning of the gene targeting strategy should consider both vector design and the processes involved in the characterization and validation of the newly formed line. The Strategic Planning section contains guidelines regarding the structure of the targeting vector. One parameter that appears to influence the frequency of homologous recombination in hESCs is the length of the homology arms (Zwaka and Thomson, 2003). Having at least one homology arm >6 kb improves the targeting frequency (Zwaka and Thomson, 2003; A.G. Elefanty and E.S. Stanley, unpub. observ.). However, the origin of the homology arms does not appear to significantly influence the targeting frequency between different hESC lines (Costa et al., 2007). The reduced requirement for the homology arms to be derived from isogenic DNA may reflect the lower frequency of polymorphisms that we have observed between different human DNA isolates compared to DNA from different inbred mouse strains (R. Davis, A.G. Elefanty, and E.G. Stanley, unpub. observ.). Therefore, we find that commercially available bacterial artificial chromosome DNA is a convenient source of genomic DNA. Similar to mouse ESCs, the frequency of homologous recombination in hESCs also appears to be locus dependent (Hasty et al., 1994; Costa et al., 2007). If homologous recombination is not obtained after the initial electroporation, the experiment should be repeated until at least 500 stably transfected clones have been screened. If targeting is still unsuccessful, we have achieved success by altering the length of the homology arms. Another alternative is to target a different region of the gene. If the generation of a targeted line in which the loss of one allele is anticipated to result in haplotype insufficiency, targeting the 3 untranslated region of the gene with a vector including an Internal Ribosomal Entry Site (IRES) upstream of the reporter coding sequences may circumvent such a problem. A fluorescent reporter marker, usually a green fluorescent (GFP) or red fluorescent (RFP) protein, is the typical reporter gene used to generate a reporter knock-in hESC line (Zwaka and Thomson, 2003; Irion et al., 2007; Davis et al., 2008a). The fluorescent protein provides an easy visual identifier of cells



corresponding to the desired population, as well as enabling the quick assessment of how the differentiating cells respond to different stimuli. In hESCs, the expression of GFP can persist beyond the timeframe that the lineagespecific marker is detected, making it also useful for lineage tracing experiments (Davis et al., 2008a). In instances where this function is not desirable, another reporter such as nonfunctional versions of cell surface receptors (e.g., CD4 or CD25) could be used, as has been done in mESCs (Yasunaga et al., 2005; Gadue et al., 2006). While cells expressing surface markers cannot be readily visualized, antibodies are available that enable the detection and isolation of these cells by immunofluorescence or flow cytometry. Alternatively, if direct visualization of the reporter knock-in cells is a requirement, then an unstable form of GFP that has a significantly reduced half-life could be utilized (Corish and Tyler-Smith, 1999). However, the cumulative fluorescence from such a GFP may also be reduced, making direct visualization of expressing cells difficult. Culturing of hESCs A homogeneous population of undifferentiated hESCs is required for these protocols. Most culturing systems consist of hESCs grown on mitotically inactivated MEFs, which provide some of the necessary factors for the maintenance and survival of hESCs in vitro. The quality of the mitotically inactivated MEFs can therefore significantly affect the hESCs culture. The hESCs should be routinely analyzed by flow cytometry for the expression of a panel of stem cell markers. Experience in passaging and maintaining undifferentiated hESCs in bulk culture is recommended before embarking on the protocols described in this unit. It is vital that the hESCs are adapted to enzymatic passaging as single cells before being transfected or cloned. Care should always be taken when enzymatically harvesting the hESCs. The dissociation treatment should last only as long as necessary to dislodge the cells from the tissue culture plastic ware. The clumps can then be gently broken up by trituration into a suspension of (predominantly) single cells. Prolonged enzymatic passaging of hESCs can select for cells adapted to this culturing technique and even result in the acquisition of chromosomal aberrations that offer a selective advantage (Draper et al., 2004). When generating genetically modified lines, the parental cell lines should be enzy-

matically passaged no more than 10 times before use. The hESCs are always passaged the day prior to an application so that the following day the cultures are semi-confluent and therefore actively proliferating before being utilized. The cells are also supplemented with fresh medium a few hours before transfecting or cloning the cells. Following the generation or cloning of gene-targeted hESC lines, the cells should be returned to organ culture dishes and maintained as dense colonies that are mechanically passaged once a week. Re-entering the organ culture phase reduces the chance of obtaining chromosomally abnormal lines. Stocks of the lines should also be frozen in liquid nitrogen. Electroporation of the targeting construct and selection of stably transfected hESCs Viable hESC colonies should emerge in culture within 2 days following electroporation (Fig. 5B.1.3B). Very high mortality levels observed following electroporation might be a consequence of excessive or rough handling of the cells throughout the protocol. Also, ensure the hESCs are resuspended in ice-cold PBS prior to electroporation and later transferred to prewarmed hESC medium (37◦ C). The electroporated cells should be washed with hESC medium before plating to remove any cellular debris that might be detrimental to the surviving hESCs. We believe that these steps improve the recovery level of the hESCs following electroporation. High concentrations of plasmid DNA can also adversely affect hESC viability and using a reduced quantity of DNA can lower mortality during electroporation, particularly if the cell count is below 1 × 107 . An endotoxin-free plasmid purification kit should be used when preparing the plasmid DNA. Rapid differentiation of the hESCs after electroporation suggests that the culture conditions may have been suboptimal or the electroporation parameters were too harsh. Under these circumstances, often the cells cannot be rescued and it is best to discard the cells and re-commence the electroporation procedure. To minimize the risk of differentiation in future transfections, the MEF density on the dishes and/or the concentration of FGF2 in the hESC medium could be increased. In addition, the hESC medium should be changed more frequently, and the dishes supplemented with MEFs earlier on during the drug selection period.



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Following selection, drug-resistant colonies should be visible within 1 week. If no colonies have formed, then the positive selection cassette in the targeting construct may be nonfunctional. Prior to electroporation, the expression of the positive selection cassette should be confirmed in another cell type, such as the human embryonic kidney (HEK) 293T line. Another possibility for the absence of hESC colonies is a failure of the electroporation procedure to transfect the vector into the cells. An abnormally short time constant (e.g., 70% confluent or are not growing as a monolayer of cells. Because of the short exposure time to puromycin, this selection method can be used even if the supporting MEF cell line is puromycin sensitive. However, the dishes must be replenished with fresh MEFs immediately post puromycin selection to support the growth of the remaining viable hESCs. Again, the initial procedure to identify the hESC clones that have excised the positive



selection marker from the targeted locus relies on PCR-based screening. Using a primer against sequences in the reporter gene in combination with a primer that matches sequences 3 of the positive selection marker, a PCR product should be amplified regardless of whether the cassette has been excised or not. If no product is obtained, this suggests that either the PCR conditions were suboptimal, or the original gene-targeted hESC line was not clonal and included nontargeted hESCs. To optimize the PCR conditions, include as a positive control a sample of DNA from the gene-targeted hESC line that still contains the positive selection marker. If the original gene-targeted hESC line is not clonal, the researcher should continue to screen colonies that were puromycin-resistant by PCR until they identify clones that are correctly targeted and have lost the positive selection cassette. These hESCs will also need to be clonally isolated using single-cell deposition flow cytometry. PCR should be performed to confirm that the Cre recombinase expression vector did not integrate into the genome. It is also recommended that the chosen hESC clones be reexposed to the selection agents to confirm their sensitivity. Generally, almost all of the colonies screened have the positive selection cassette excised (Davis et al., 2008b). Cloning It is important that all reporter knock-in hESC lines that will be used as reagents in future experiments are clonally isolated. This ensures that the final cell population is homogeneous with respect to the targeted locus and the excision of the antibiotic-selection cassette. For example, a mixed population of targeted and nontargeted hESCs could result in the misclassification of cell types if identification is based on the expression of the reporter gene. The cloning efficiency of hESCs using single-cell deposition flow cytometry varies between cell lines. Always increase the concentration of FGF2 in the hESC medium to 40 ng/ml during cloning. This helps to grow and expand colonies from individual hESCs and reduces the level of spontaneous differentiation (Amit et al., 2000; Xu et al., 2005). If the colonies still spontaneously differentiate on the 96-well plates, increase the frequency of hESC medium changes and/or the density of the mitotically inactivated MEF cells in the wells.

If less than 10 viable hESC colonies are obtained following the isolation of ∼1000 individual hESCs, 2 μM of the Rho-associated kinase inhibitor, Y-27632, could be added to the cells following single-cell deposition. This could improve the level of survival of the hESCs; however, the poor cloning efficiency could also be due to the ratio of MEFs to hESCs being too high in the sorted population. Reducing the concentration of feeder cells by passaging the hESCs onto a lower density of MEFs (∼1 × 104 /cm2 ) the day before singlecell cloning reduces the background feeder count. On the day of single-cell deposition, the flasks should be ∼80% confluent with hESCs.

Anticipated Results In most cases, if the guidelines and protocols described in this unit are adhered to, a reporter knock-in hESC line will be generated in which the expression of the reporter accurately reflects that of the targeted gene. The hESCs will also retain their stem cell characteristics and remain karyotypically normal. At the time of writing, the authors’ laboratory has used these protocols to generate reporter knock-in hESC lines at nine different loci, with multiple targeted clones in two independent hESC lines for most cases. The electroporation protocol (Basic Protocol 1) should yield stably transfected hESC clones at a frequency of 20 to 500 per 107 input hESCs. However, the frequency of homologous recombination amongst these surviving clones will vary depending on the vector used and the genomic locus being targeted. In the authors’ laboratory, this frequency has ranged from 1.3% to 14.6%. The method for the removal of loxPflanked positive selection cassettes from the genome of genetically modified hESC lines (Basic Protocol 2) has proven to be very successful. On average, 62 puromycin-resistant colonies are obtained per 1 × 106 hESCs transfected with the pEFBOS-creIRESpuro expression vector, with >95% of the resulting colonies screened having the selection cassette removed. The cloning efficiency of the hESCs using single-cell deposition flow cytometry is generally between 4% and 8%.

Time Considerations Once the targeting vector has been constructed, it will take a minimum of 6 months to generate and validate a reporter knock-in hESC line. However, it could take several more



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months if a correctly targeted hESC clone is not obtained from the first electroporation attempt. On average, it takes 1 year from designing and constructing the targeting vector to having an established and proven targeted reporter knock-in hESC line. The electroporation and subsequent selection of hESC clones that have integrated the targeting construct takes ∼2 1/2 weeks. The expansion of these clones and isolation of DNA for PCR screening requires a further 2 weeks of culturing. This can be reduced by ∼1 week if DNA is prepared directly from the hESC clones on the “Primary” 48-well plates, prior to replicating plates. The PCR screen should take no more than 2 days. Because this entire procedure takes ∼1 month, it is recommended that several consecutive electroporations be initiated at weekly intervals, even before the PCR screening results from the first transfection are known. Once a correctly targeted hESC clone is obtained, it will take at least 4 weeks of culturing to expand the clone for removal of the positive selection cassette. Additionally, these hESC lines should be cryopreserved and karyotyped, and homologous recombination confirmed by Southern blot analysis. The procedure for the removal of the positive selection cassette and subsequent confirmation by PCR will also take ∼1 month. The transfection of the pEFBOS-creIRESpuro vector and selection by puromycin requires 5 days. The transfected hESCs then require at least another 10 days before they can be transferred to 48-well plates for expansion. After the expansion of these clones (∼2 weeks), the extraction of DNA and PCR analysis will take 3 days. The expansion of two or three hESC clones in which the selection cassette has been removed will take another month. The cloning of these lines by single-cell deposition flow cytometry and confirmation that the cells contain the correct genetic modification requires a further 4 weeks. If the cloning protocol is integrated into the procedure for removing the positive selection cassette, it is possible to reduce the length of time required to generate a reporter knock-in hESC line by ∼2 months.

Council (NHMRC) of Australia. AGE is a Senior Research Fellow of the NHMRC.

Acknowledgements

Eiges, R., Schuldiner, M., Drukker, M., Yanuka, O., Itskovitz-Eldor, J., and Benvenisty, N. 2001. Establishment of human embryonic stem celltransfected clones carrying a marker for undifferentiated cells. Curr. Biol. 11:514-518.

We thank Robyn Mayberry, Kathy Koutsis, and Amana Bruce for the provision of hESCs. This work was supported by the Australian Stem Cell Centre (ASCC), the Juvenile Diabetes Research Foundation (JDRF), and the National Health and Medical Research

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Guidelines for the Conduct of Human Embryonic Stem Cell Research

APPENDIX 1A

ABSTRACT This appendix provides a summary of and links to the ISSCR Guidelines for the Conduct of Human Embryonic Stem Cell Research and supporting documents. Curr. Protoc. Stem C 2009 by John Wiley & Sons, Inc. Cell Biol. 9:A.1A.1-A.1A.1. Keywords: embryonic stem cells r guidelines r ISSCR research

The International Society for Stem Cell Research (ISSCR) calls for due consideration and appropriate oversight of human embryonic stem cell research to ensure transparent ethical and responsible performance of scientific experiments. These Guidelines, prepared by an ISSCR Task Force composed of international representatives, are meant to emphasize the responsibility of scientists to ensure that human embryonic stem cell research is carried out according to rigorous standards of research ethics, and to encourage uniform research practices that should be followed by all human stem cell scientists globally. The scope of these Guidelines includes principles for review and approval as well as oversight, enforcement mechanisms, procurement of materials with informed consent, principles for derivation, banking, and distribution of stem cell lines, dispute resolution, and ongoing review. There is an appendix with a glossary of scientific terms and a set of links to local, national, and international regulations related to embryonic stem cell research. These Guidelines are published at: http://www.isscr.org/guidelines/index.htm with links to Sample Consent documents for procuring materials, a sample Material Transfer Agreement (MTA), a Science magazine policy forum summarizing the guidelines, and biographies of the task force members.

Useful Information Current Protocols in Stem Cell Biology A.1A.1 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sca01as9 C 2009 John Wiley & Sons, Inc. Copyright

A.1A.1 Supplement 9

ISSCR Guidelines for the Clinical Translation of Stem Cells

APPENDIX 1B

ABSTRACT This appendix provides a summary of and links to the ISSCR Guidelines for the Clinical Translation of Stem Cells and supporting documents. Curr. Protoc. Stem Cell Biol. C 2009 by John Wiley & Sons, Inc. 9:A.1B.1-A.1B.1. Keywords: stem cells r guidelines r ISSCR clinical translation

In the fast-paced world of stem cell research, with the ever-present pressures for positive results that will translate to therapeutic use of human stem cells and their products, there is need for guidance and oversight to ensure that the scientific, clinical, regulatory, and social issues are properly addressed to guarantee that research results are appropriately translated into clinical therapies. These Guidelines define a roadmap for researchers and clinicians, outlining what needs to be accomplished to move stem cells from promising research to proven treatments for patients. They address issues related to cell processing and manufacture, preclinical studies, and clinical research, and contain 40 recommendations to ensure scientifically based, transparent, ethical, and medically and socially responsible clinical translation of stem cell–based therapies. Additionally, these Guidelines contain four appendices: the first is a Patient Handbook for those considering stem cell therapy; the second is a list of sites for local, national, and international regulations; the third is a list of related articles; and the fourth is a set of definitions. The Guidelines can be accessed at: http://www.isscr.org/clinical trans/index.cfm with links to the Patient Handbook (Appendix 1), additional resources (Appendix 2), a summary article from Cell Stem Cell, and the ISSCR and Cell Stem Cell press release.

Useful Information Current Protocols in Stem Cell Biology A.1B.1 Published online April 2009 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/9780470151808.sca01bs9 C 2009 John Wiley & Sons, Inc. Copyright

A.1B.1 Supplement 9

Standard Laboratory Equipment

APPENDIX 2

Listed below are pieces of equipment that are standard in the modern stem cell biology laboratory—i.e., items used extensively in this manual and thus not usually included in the individual materials lists. No attempt has been made to list all items required for each procedure in the Materials list of each protocol; rather, those lists note those items that might not be readily available in the laboratory or that require special preparation. See SUPPLIERS APPENDIX for contact information for commercial vendors of laboratory equipment. Applicator, cotton-tipped and wooden Autoclave Bag sealer Balances, analytical and preparative Beakers Bench protectors, plastic-backed (including “blue” pads) Biohazard disposal containers and bags Biosafety cabinet, tissue culture or laminar flow hood; filters air and maintains air flow pattern to protect cultured cells from investigator and vice versa Bottles, glass, plastic, and squirt Bunsen burners Centrifuges, low-speed (to 20,000 rpm) refrigerated, ultracentrifuge (20,000 to 80,000 rpm), large-capacity low-speed, tabletop, with appropriate rotors and adapters Centrifuge tubes and bottles, plastic and glass, various sizes Clamps Conical centrifuge tubes, plastic and glass Containers, assortment of glass and plastic, for gel and membrane washes Coplin jars, glass, for 25 × 75–mm slides Cryovials, sterile (e.g., Nunc) Cuvettes Desiccator and desiccant Dry ice Electrophoresis equipment, agarose and acrylamide, full-size and mini, with power supplies Film developing system and darkroom Filtration apparatus Forceps Fraction collector Freezers, −20◦ C, −70◦ C, and liquid nitrogen Fume hood Geiger counter Gel dryer Gloves, disposable plastic and heat resistant Graduated cylinders Heating blocks, thermostatically controlled for test tubes and microcentrifuge tubes

Hemacytometer and/or electronic cell counter Homogenizer Humidified CO2 incubator Ice bucket Ice maker Immersion oil for microscopy Lab coats Laboratory glassware Light box Liquid nitrogen Lyophilizer Magnetic stirrer, with and without heater, and stir bars Markers, including indelible markers, china-marking pens, and luminescent markers Microcentrifuge, Eppendorf-type with 12,000 to 14,000 rpm maximum speed Microcentrifuge tubes, 0.2-, 0.5-, 1.5-, 2-ml Microscope slides, glass, 25 × 75–mm, and coverslips Microscope with camera, upright, inverted, fluorescence, phase-contrast, dissecting Microtiter plate reader Mortar and pestle Ovens, drying and microwave Paper cutter, large Paper towels Parafilm Pasteur pipets and bulbs PCR thermal cycler and tubes pH meter pH paper Pipets, graduated Pipettors, adjustable delivery, 0.5- to 10-µl, 10- to 200-µl, and 200- to 1000-µl Polaroid camera or video documentation system Power supplies, 300-V for polyacrylamide gels, 2000- to 3000-V for other applications Racks, test tube and microcentrifuge tubes Radiation shield, Lucite or Plexiglas Laboratory Equipment

Current Protocols in Stem Cell Biology (2007) A.2.1-A.2.2 C 2007 by John Wiley & Sons, Inc. Copyright

A.2.1 Supplement 1

Radioactive waste containers for liquid and solid wastes Refrigerator, 4◦ C Ring stand and rings Rubber policemen or plastic scrapers Rubber stoppers Safety glasses Scalpels and blades Scintillation counter, β Scissors Shakers, orbital and platform, room temperature or 37◦ C Spectrophotometer, visible and UV range Speedvac evaporator Syringes and needles Tape, masking, electrician’s black, autoclave, and Time tape Test tubes, glass and plastic, various sizes, with and without caps

Timer Toolbox with common tools Trays, plastic and glass, various sizes Tubing, rubber and Tygon UV light sources, long- and short-wavelength UV transilluminator UV transparent plastic wrap (e.g., Saran Wrap) Vacuum desiccator Vacuum oven Vacuum supply Vortex mixers Waring blendor Water bath with adjustable temperature Water purification system X-ray film cassettes and intensifying screens

Standard Laboratory Equipment

A.2.2 Supplement 1


SELECTED SUPPLIERS OF REAGENTS AND EQUIPMENT Listed below is contact information for commercial suppliers who have been recommended for particular items used in Current Protocols in Stem Cell Biology because: (1) the particular brand has actually been found to be of superior quality, or (2) the item is difficult to find in the marketplace. Consequently, this compilation may not include some important vendors of biological supplies. For comprehensive listings, see Linscott’s Directory of Immunological and Biological Reagents (Santa Rosa, CA), The Biotechnology Directory (Stockton Press, New York), the annual Buyers’ Guide supplement to the journal Bio/Technology, as well as various sites on the Internet.

Abcam 617-225-2272 Fax: 617-507-5831 http://www.abcam.com

Applied Biosystems 650-638-5800 Fax: 650-638-5998 http://www.appliedbiosystems.com

Bio-Rad Laboratories 800-424-6723 Fax: 510-741-1000 http://www.bio-rad.com

Curis 617-503-6500 Fax: 617-503-6501 http://www.curis.com

Addgene http://www.addgene.org

Applied Imaging 800-634-3622 Fax: 408-562-0250 http://www.aicorp.com

Calbiochem-Novabiochem 800-854-3417 Fax: 800-776-0999 858-450-9600 http://www.calbiochem.com

Dako Denmark A/S 45-44-85-95-00 http://www.dako.com

Affymetrix 888-DNA-CHIP (888-362-2447) Fax: 408-731-5441 http://www.affymetrix.com Agar Scientific 44-1279-813-519 Fax: 44-1279-815-106 http://www.agarscientific.com Agilent Technologies 877-424-4536 408-345-8886 Fax: 408-345-8474 http://www.home.agilent.com Amaxa See Lonza Ambion 512-651-0200 http://www.ambion.com American Pharmaceutical Partners 847-969-2700 888-391-6300 Fax: 800-743-7082 http://www.appdrugs.com American Type Culture Collection (ATCC) 800-638-6597 Fax: 703-365-2700 http://www.atcc.org Amersham Bioscience 732-457-8000 http://www.amersham.com Amgen 805-447-1000 Fax: 805-447-1010 http://www.amgen.com Amresco 800-829-2805 Fax: 440-349-1182 440-349-1199 http://www.amresco-inc.com/

Arctur See Molecular Devices Corporation ATCC See American Type Culture Collection Bando Chemical Industries http://www.bandousa.com Bard Parker See Becton Dickinson Baxter Healthcare 800-777-2298 Fax: 847-948-2000 http://www.baxter.com BD http://www.bd.com BD Biosciences 877-232-8995 http://bdbiosciences.com BD Falcon See BD Biosciences BD Pharmingen 800-848-6227 Fax: 858-812-8888 858-812-8800 http://www.pharmingen.com Beckman Coulter 800-742-2345 Fax: 800-643-4366 http://www.beckmancoulter.com Becton Dickinson Labware 888-237-2762 Fax: 800-847-2220 201-847-4222 http://www.bdfacs.com Bioline 888-257-5155 Fax: 775-828-0202 http://www.bioline.com Biomeda 800-341-8787 Fax: 650-341-8787 http://www.biomeda.com

Cambrex Corporation 201-804-3000 http://www.cambrex.com Canemco Inc. & Marivac Inc. 450-562-1451 Fax: 450-562-3430 http://www.canemco.com Cellomics 800-432-4091 412-770-2500 Fax: 412-770-2450 http://www.cellomics.com Chemicon International 800-437-7500 Fax: 909-676-8080 http://www.chemicon.com Clontech Laboratories 800-662-2566 Fax: 650-424-8222 http://www.clontech.com Cole-Parmer Instrument 800-323-4340 Fax: 847-247-2929 847-549-7600 http://www.coleparmer.com

Developmental Studies Hybridoma Bank 319-335-3826 http://dshb.biology.uiowa.edu/ DiaMed Lab Supplies Inc. 800-434-2633 Fax: 905-625-6280 http://www.diamed.ca Dow Corning 989-496-4400 Fax: 989-496-6731 http://www.dowcorning.com eBioscience 858-642-2058 Fax: 858-642-2046 http://www.ebioscience.com Eppendorf 800-645-3050 516-334-7500 Fax: 516-334-7506 http://www.eppendorf.com Ethicon See Johnson & Johnson Health Care Systems Falcon See Becton Dickinson Labware

Cooper Medical 64-9-3022955 http://www.coopermedical.co.nz

Fine Science Tools 800-521-2109 Fax: 650-349-1636 http://www.finescience.com

Corning and Corning Science Products 800-222-7740 Fax: 607-974-9000 http://www.corning.com

Fisher Scientific 800-766-7000 Fax: 412-562-8300 http://www3.fishersci.com

Costar See Corning Crown Scientific 61-1300-727-696 Fax: 61-1300-135-123 http://www.crownscientific.com.au

Fluka Chemical See Sigma-Aldrich GE Healthcare http://www.gehealthcare.com Gelaire http://www.gelaire.com.au

Suppliers

1 Current Protocols in Stem Cell Biology

CPSC Supplement 8

Gemini BioProducts 818-591-3530 http://www.gembio.com/ Gibco See Invitrogen Global Stem 888-545-0238, ext. 114 301-545-0238, ext. 114 Fax: 301-424-1989 http://www.globalstem.com Greiner Bio-One http://www.greinerbioone.com Hamilton Thorne Biosciences http://www.hamiltonthorne.com Hausser Scientific 215-675-7769 Fax: 215-672-9602 http://www.hausserscientific.com

Johnson & Johnson Health Care Systems 800-255-2500 Fax: 732-562-2212 http://www.ecatalog.ethicon.com Kendro Laboratory Products 800-522-SPIN Fax: 203-270-2166 203-270-2080 http://www.kendro.com Kent Scientific 888-572-8887 Fax: 860-567-4201 860-567-5496 http://www.kentscientific.com Kord-Valmark 800-452-9070 http://www.kord-valmark.com

Hunter Antisera 61-49620967 http://www.hum-molgen.org

Lafayette Instrument 800-428-7545 Fax: 765-423-4111 765-423-1505 http://www.lafayetteinstrument.com

HyClone Laboratories 800-HYCLONE Fax: 801-753-4584 http://www.hyclone.com

LEC Instruments 61-3-9763-0080 Fax: 61-3-9764-0086 http://lecinstruments.com

Illumina 800-809-4566 858-202-4566 Fax: 858-202-4766 http://www.illumina.com

Lonza http://www.lonza.com

Instrumedics 314-522-8671 Fax: 314-522-6360 http://www.instrumedics.com Integrated DNA Technologies 800-328-2661 319-626-8400 Fax: 319-626-8444 http://www.idtdna.com Invitrogen 800-955-6288 Fax: 760-603-7200 http://www.invitrogen.com Irvine Scientific 800-577-6097 Fax: 949-261-7800 http://www.irvinesci.com ISC BioExpress 800-999-2901 Fax: 801-547-5047 http://www.bioexpress.com The Jackson Laboratory 207-288-6000 http://www.jax.org John Morris Scientific http://www.johnmorris.com.au

Marienfeld 49-0-9343-6272-0 Fax: 49-0-9343-6272-25 http://www.superior.de/

MJ Research 800-PELTIER Fax: 617-923-8000 http://www.mjr.com

Qiagen 800-426-8157 Fax: 800-718-2056 http://www.qiagen.com

Molecular BioProducts 800-995-2787 Fax: 858-453-7551 http://www.mbpinc.com

Quality Biological 800-443-9331 301-840-8950 Fax: 301-840-0743 http://www.qualitybiological.com

Molecular Devices 408-548-6000 http://www.moldev.com Molecular Probes 800-438-2209 Fax: 541-465-8300 http://www.probes.com MP Biomedicals 800-854-0530 Fax: 800-334-6999 http://www.mpbio.com Nalge Nunc International 800-625-4327 Fax: 716-264-9346 http://www.nalgenunc.com Novocastra 44-0-191-215-0567 Fax: 44-0-191-215-1152 http://www.ebiotrade.com/buyf/ Novocastra/index.htm Orange Scientific 32-2-387-56-97 http://www.orangesci.com Pall http://www.pall.com

Matrix Technologies Corp. 800-345-2060 Fax: 603-595-0106 http://www.matrixtechcorp.com

Partec 49-2534-8008-0 Fax: 49-2535-8008-90 http://www.partec.com

MDS Nordion 800-465-3666 Fax: 613-592-6937 613-592-2790 http://www.mds.nordion.com

PeproTech 800-436-9910 Fax: 609-497-0253 http://www.peprotech.com

Mediatech/Cellgro 800-CELLGRO Fax: 703-471-0363 http://www.cellgro.com MidAtlantic Diagnostics 800-648-1151 Fax: 856-762-2000 http://www.midatlanticdiagnostics.com Millipore 800-645-5476 Fax: 800-645-5439 http://www.millipore.com Miltenyi Biotec 800-367-6227 Fax: 530-888-8871 http://www.miltenyibiotec.com

PerkinElmer 800-762-4000 http://www.perkinelmer.com

R & D Systems 800-343-7475 Fax: 612-379-2956 http://www.rndsystems.com Research Diagnostics 800-631-9384 Fax: 973-584-0210 973-584-7093 http://www.researchd.com Research Products International 800-323-9814 Fax: 847-635-1177 847-635-7330 http://www.rpicorp.com Roche Applied Science 800-262-1640 Fax: 800-428-2883 https://www.roche-applied-science.com Roche Diagnostics 800-262-1640 Fax: 317-845-2000 http://www.roche.com Sakura 800-725-8723 310-972-7800 Fax: 310-972-7888 http://www.sakuraus.com Santa Cruz Biotechnology 800-457-3801 Fax: 831-457-3800 http://www.scbt.com Henry Schein 631-843-5500 http://www.henryschein.com

Phoenix Biomedical Products 905-670-8299 Fax: 905-670-0195 http://www.phoenix-biomed.com

Serotec 44-1865-852722 Fax: 44-1865-373899 In the US: 800-265-7376 http://www.serotec.co.uk

Nicholas Piramal 91-22-3046-6666 http://www.nicholaspiramal.com

Sigma-Aldrich 800-358-5287 http://www.sigma-aldrich.com

Polysciences 800-523-2575 http://www.polysciences.com

Sorvall See Kendro Laboratory Products

Promega 800-356-9526 Fax: 608-274-4330 http://www.promega.com

Southern Biotechnology Associates 800-722-2255 Fax: 205-945-8768 205-945-1774 http://southernbiotech.com

Suppliers

2 CPSC Supplement 8


Specialty Media 800-543-6029 Fax: 908-454-7774 http://www.specialtymedia.com

Taconic 888-TACONIC (888-822-6642) 518-697-3915 http://www.taconic.com

Thermo Electron 732-627-0220

Vectashield See Vector Laboratories

Thermo Scientific http://www.thermo.com

SpeedVac http://www.thermo.com

Techne 800-225-9243 Fax: 609-987-8177 609-452-9275 http://www.techneusa.com

Tree Star Software 800-366-6045 http://www.treestar.com

Vector Laboratories 800-227-6666 Fax: 650-697-3600 http://www.vectorlabs.com

Stem Cell Technologies 800-667-0322 Fax: 604-877-0713 http://www.stemcell.com Sterilin 44-0-844-844-3737 Fax: 44-0-844-844-2373 http://www.sterilin.co.uk Stoelting 920-894-2293 Fax: 920-894-7029 http://www.stoelting.com C. L. Sturkey 717-274-9441 Fax: 717-274-9442 http://www.sturkey.com Swemed See Vitrolife

Techno Plastic Products http://www.tpp.ch/ Ted Pella 800-237-3526 Fax: 530-243-2200 http://www.tedpella.com Tel-Test 281-482-2672 Fax: 281-482-1070 http://www.bioresearchonline.com/ storefronts/teltest.html Terumo Medical 800-283-7866 Fax: 732-302-3083 732-302-4900 http://www.terumomedical.com

Tyco Healthcare http://www.tyco.com United States Biological (US Biological) 800-520-3011 Fax: 781-639-1768 http://www.usbio.net Univentor 356-21-895824 Fax: 356-21-895835 www.univentor.com USA Scientific 800-LAB-TIPS Fax: 352-351-2057 352-237-6288 http://www.usascientific.com VacSax http://www.vacsax.plus.com

Vision BioSystems 800-753-7264 (North America) Fax: 781-610-1190 http://www.vision-bio.com Vitrolife 46 31 721 80 00 http://www.vitrolife.com VWR 800-932-5000 Fax: 610-431-9174 http://www.vwrsp.com Whatman 800-WHATMAN 973-245-8300 Fax: 973-245-8324 http://www.whatman.com Worthington Biochemical 800-445-9603 Fax: 732-462-3838 http://www.worthington-biochem.com

Suppliers

3 Current Protocols in Stem Cell Biology

CPSC Supplement 8

Current Protocols in Stem Cell Biology

Hematopoietic Stem Cell Protocols

Hematopoietic Stem Cell Protocols

Stem Cell Biology

Hematopoietic Stem Cell Biology

Hematopoietic Stem Cell Biology (Stem Cell Biology and Regenerative Medicine)

Human Embryonic Stem Cell Protocols

Human Embryonic Stem Cell Protocols (Methods in Molecular Biology)

Human Embryonic Stem Cell Protocols (Methods in Molecular Biology)

Human Embryonic Stem Cell Protocols

Current Protocols in Molecular Biology

Current Protocols in Molecular Biology

Current Protocols in Molecular Biology

Cell Biology Protocols

Current protocols Molecular Biology

Essentials of Stem Cell Biology

Stem Cell Biology in Health and Disease

Trends in Stem Cell Biology and Technology

Advances in Cancer Stem Cell Biology

Trends in Stem Cell Biology and Technology

Stem Cell Assays (Methods in Molecular Biology)

Allogeneic Stem Cell Transplantation (Current Clinical Oncology)

Embryonic Stem Cell Protocols. Differentiation Models

Embryonic Stem Cell Protocols: Isolation and Characterization

Embryonic Stem Cell Protocols. Isolation and Characterization

B Cell Protocols (Methods in Molecular Biology)

Current Protocols in Molecular Biology (1988 - 2003)

Current Protocols in Mouse Biology (Volume 1)

B Cell Protocols (Methods in Molecular Biology)

Essentials of Stem Cell Biology, Second Edition

Stem Cell Biology and Gene Therapy

Current Protocols in Stem Cell Biology

Hematopoietic Stem Cell Protocols

Hematopoietic Stem Cell Protocols

Stem Cell Biology

Hematopoietic Stem Cell Biology

Hematopoietic Stem Cell Biology (Stem Cell Biology and Regenerative Medicine)

Human Embryonic Stem Cell Protocols

Human Embryonic Stem Cell Protocols (Methods in Molecular Biology)

Human Embryonic Stem Cell Protocols (Methods in Molecular Biology)

Human Embryonic Stem Cell Protocols

Current Protocols in Molecular Biology

Current Protocols in Molecular Biology

Current Protocols in Molecular Biology

Cell Biology Protocols

Current protocols Molecular Biology

Essentials of Stem Cell Biology

Stem Cell Biology in Health and Disease

Trends in Stem Cell Biology and Technology

Advances in Cancer Stem Cell Biology

Trends in Stem Cell Biology and Technology

Stem Cell Assays (Methods in Molecular Biology)

Allogeneic Stem Cell Transplantation (Current Clinical Oncology)

Embryonic Stem Cell Protocols. Differentiation Models

Embryonic Stem Cell Protocols: Isolation and Characterization

Embryonic Stem Cell Protocols. Isolation and Characterization

B Cell Protocols (Methods in Molecular Biology)

Current Protocols in Molecular Biology (1988 - 2003)

Current Protocols in Mouse Biology (Volume 1)

B Cell Protocols (Methods in Molecular Biology)

Essentials of Stem Cell Biology, Second Edition

Stem Cell Biology and Gene Therapy

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